Integrated Pest Management for Christmas Tree Production: A ...

IN MEMORIAM 

We dedicate this publication to Dr. Paul Heller and 

Dr. William Merrill for their contribution to the 

Christmas Tree IPM Program and for many years of 

commitment to helping growers in Pennsylvania 

and throughout the United States. 

Paul R. Heller (1948–2010) 

Professor of Entomology 

The Pennsylvania State University 

1976–2010 

William Merrill Jr. (1933–2003) 

Professor Emeritus of Plant Pathology 


1965–1999

PREFACE 

The Pennsylvania Integrated Pest Management 

Program (PA IPM) is pleased to provide Integrated Pest 

Management for Christmas Tree Production, designed 

to help Christmas tree growers manage pests based 

on sound integrated pest management principles. PA 

IPM is a collaboration of the Pennsylvania Department 

of Agriculture (PDA) and The Pennsylvania State 

University. This partnership has resulted in excellent 

service to Pennsylvania growers as well as to growers 

in surrounding states. The institutional collaboration 

embodied by PA IPM allows us to draw on Department 

of Agriculture technical experts and Penn State faculty 

and extension educators. 

This manual is part of an educational program that 

was initiated in 1985 by Dr. William Merrill Jr., professor 

of plant pathology, for the thriving Pennsylvania Christmas 

tree industry. Dr. William Merrill and Ms. Nancy 

Wenner, along with other personnel from the Departments 

of Plant Pathology, Horticulture, and Entomology, 

organized the fi rst Christmas Tree Pest Management 

Short Course held at Penn State University Park. Over 

the years, the course has evolved but is still held each 

winter in State College to provide growers with handson 

experiences in diagnosing the common insect and 

disease problems on Christmas trees. 

In 1991, Penn State entomologist Dr. Paul Heller 

initiated the Christmas Tree Landscape Pest Management 

Newsletter. The newsletter was mailed to county 

extension offi ces and interested growers on a regular 

basis. PDA entomologists Rayanne Lehman and Dr. Karl 

Valley began contributing scouting information to the 

newsletter in 1992. PDA staff continued the scouting 

program with support from PDA by providing weekly 

scouting reports for growers. The scouting report is 

still produced weekly during the growing season with 

support from PDA and growers. 

To enhance this educational outreach, and in 

response to the needs of Christmas tree growers for 

pest control information pertinent to Pennsylvania, a 

committee comprising scientists, regulators, extension 

educators, and growers was assembled to discuss 

their needs. One need was a resource customized for 

Pennsylvania conditions that describes pests and their 

management. An advisory body was assembled to help 

design and review the manual, and several experts 

contributed text and photos. 

We hope this manual will encourage growers 

to develop their own IPM programs and train their 

employees to become familiar with IPM practices. 

Funding and support to write and review this 

manual were provided by the Pennsylvania Department 

of Agriculture, The Pennsylvania State University, 

and through grant support from the Environmental 

Protection Agency Region 3. 

—Cathy Thomas, IPM Coordinator, 

Pennsylvania Department of Agriculture 

—Ed Rajotte, Professor of Entomology and 

Penn State IPM Coordinator 

HOW TO USE THIS MANUAL 

The purpose of this manual is to help growers identify, 

monitor, and control disease and insect pests affecting 

Christmas trees. The pests included are common on 

Christmas trees in the Mid-Atlantic and Northeast 

regions of the United States; however, many of these 

pests can be found in other regions of the United 

States. This manual is intended for fi eld use by any level 

of grower, whether professional or hobbyist. Unfamiliar 

terms are defi ned in the glossary at the back of the 

manual. 

Steps for using this manual to identify a pest 

problem: 

1. Identify the tree species. 

2. Refer to Appendix A: Pest and Disease Photo 

Chart. This chart has a series of thumbnail photos 

corresponding to pests or abiotic problems. Select 

the photo(s) that best matches symptomatic plants 

and refer to the appropriate “fact sheet(s)” for additional 

information. 

Each pest fact sheet provides useful information, 

including hosts affected, damage potential, 

symptoms and signs, similar symptoms, calendar 

of activities, identifi cation tips, biology and life 

cycle, monitoring and management strategies, 

control options, and next crop/prevention strategies. 

Photographs contained in the fact sheets show 

the symptoms and life stages of the pests as seen 

throughout the growing season. 

Seasonal calendars have been included to serve 

as guides for scouting and controlling pests; growing 

degree days have been included when available. 

However, the best method for detection and control is 

still personal observation. Guidelines for scouting can 

be found on page 9 of this manual. Additionally, several 

templates (Growing Degree Day Record, Scouting Record, 

and Pesticide Record) are located in the appendix. 

These templates can be copied to help with yearly 

record keeping. 

Christmas tree scouting updates can be found at 

ento.psu.edu/extension/christmas-trees/scoutingreports 

starting in March and continuing weekly 

through June. Additional reports are included during 

the month of August. Past reports starting in 2000 are 

available for historical perspective. 

INTEGRATED 

PEST 

MANAGEMENT 

for 

CHRISTMAS 

TREE 

PRODUCTION 

A GUIDE FOR 

PENNSYLVANIA 

GROWERS

ACKNOWLEDGMENTS 

Project Coordinators 

Cathy E. Thomas, PA IPM Coordinator 


Sarah Pickel, Education Specialist 


Contributors 

Amber Brunskill, IPM Curriculum Development Assistant 

Penn State Department of Entomology 

Rayanne D. Lehman, Entomologist (Retired) 


Karen Najda, Plant Inspector, Region VII 


Tracey N. Olson, Plant Pathologist 




Brian Schildt, IPM Scouting Consultant 




Reviewers 

Steve Derstine, Farm Manager 

J. C. Hill Tree Farms, Inc. 

Jim Fogarty 

Halabura Tree Farm 

Rayanne D. Lehman, Entomologist (Retired) 


Kimberly J. Miller, Plant Inspector, Region III 


Karen Najda, Plant Inspector, Region VII 


Tracey N. Olson, Plant Pathologist 


Ed Rajotte, Professor of Entomology and 

PA IPM Coordinator 


Linda L. Signarovitz, Plant Inspector (Retired) 


Advisers 

Timothy M. Abbey, Extension Educator 

Penn State Cooperative Extension, Capital Area 

Ricky Bates, Professor of Horticulture 

Penn State Department of Horticulture 

Sandy Gardosik, Entomologist 


Ed Rajotte, Professor of Entomology and PA IPM 

Coordinator 


Linda L. Signarovitz, Plant Inspector (Retired) 


A special thanks to: 

George Perry, Extension Educator (Retired), Penn State 

Cooperative Extension, Schuylkill and Berks Counties, 

for his many years of dedicated service to Pennsylvania 

Christmas tree growers 

PA IPM Program Staff 

Ed Rajotte, Professor of Entomology and PA IPM 

Coordinator 




Kristie Auman-Bauer, PA IPM Public Relations and 

Outreach Coordinator 


David Biddinger, Tree Fruit Entomologist and 

Biocontrol Specialist 


Amber Brunskill, Curriculum Development Assistant 


Wade Esbenshade, IPM and Biocontrol Specialist 


Lyn Garling, PA IPM Manager of Programs 


Rhonda Griffi n, IPM Home Healthcare Educator 

Penn State Cooperative Extension in Philadelphia 

County 

Dion Lerman, Environmental Health Programs Specialist 


County 

Cathy Nardozzo, PA IPM Webmaster 


Michelle Niedermeier, Community IPM and 

Environmental Health Program Coordinator 


County 



Brian Schildt, IPM Scouting Consultant 

Pennsylvania Department of Agriculture

CONTENTS 

Integrated Pest Management Basics ................ 5 

Step 1. Preparing for IPM: Planning and 

Prevention ....................................................................6 

Step 2. Identifi cation and Understanding of 

Pests and Problems ......................................................7 

Insects and Mites ................................................7 

Diseases (Fungi and Nematodes) ........................8 

Vertebrates (Mammals and Birds) .......................8 

Weeds .................................................................8 

Environmental Factors .........................................8 

Step 3. Monitoring Trees for Pest Populations ..............9 

Growing Degree Days/Temperature .....................9 

Soil Temperature................................................10 

Personal Monitoring Kit ....................................10 

Monitoring Tips .................................................11 

Trapping ...........................................................11 

Diagnosis ..........................................................11 

Step 4. Setting an Action Threshold ............................12 

Step 5. Selecting Control Options ...............................12 

Cultural and Mechanical Control .......................12 

Biological Control ..............................................13 

Chemical Control ...............................................13 

Step 6. Evaluating Results of Control Actions .............16 

Pennsylvania Conifer Certifi cation and 

Quarantine Guidelines ...............................................16 

Growing Degree Day Ranges for 

Christmas Tree Pests ...................................................18 

Insect and Mite Calendar ..........................................19 

Pest Fact Sheets ................................................ 21 

Needle Discoloration and Injury .................................22 

Bagworm...........................................................23 

Balsam Twig Aphid ............................................26 

Cooley Spruce Gall Adelgid on Douglas-Fir ......29 

Cryptomeria Scale .............................................31 

Cyclaneusma Needle Cast .................................34 

Douglas-Fir Needle Midge .................................37 

Elongate Hemlock Scale ....................................39 

Eriophyid Rust and Sheath Mites ......................42 

Gypsy Moth .......................................................45 

Lophodermium Needle Cast ..............................48 

Pine Needle Scale ..............................................51 

Pine Sawfl ies .....................................................53 

Ploioderma Needle Cast ....................................56 

Red-band (Dothistroma) Needle Blight .............58 

Rhabdocline Needle Cast ..................................60 

Rhizosphaera Needle Cast.................................63 

Spruce Needle Rust ...........................................65 

Spruce Spider Mite ............................................67 

Swiss Needle Cast .............................................70 

Shoot and Branch Injury .............................................72 

Atropellis Canker ...............................................73 

Balsam Woolly Adelgid ......................................75 

Botrytis Blight ...................................................78 

Cooley Spruce Gall Adelgid on Spruce ...............80 

Diplodia (Sphaeropsis) Tip Blight of Pines and 

Other Conifers ...................................................82 

Eastern Pine Weevil ...........................................84 

Eastern Spruce Gall Adelgid ..............................86 

Gall Rusts ..........................................................88 

Pales Weevil ......................................................90 

Pine Bark Adelgid ..............................................92 

Pine Shoot Beetle ..............................................94 

Striped Pine Scale ..............................................97 

White Pine Blister Rust ......................................99 

White Pine Weevil ...........................................101 

Stem and Root Injury/Tree Mortality ........................106 

Armillaria Root Rot .........................................107 

Bark and Engraver Beetles ..............................109 

Phytophthora Root Rot....................................111 

Pine Root Collar Weevil ...................................113 

Pine Wilt Disease .............................................116 

White Grubs ....................................................118 

Zimmerman Pine Moth....................................120 

Other Types of Damage to Conifers ..........................122 

Chemical Damage ...........................................123 

Environmental Damage ...................................124 

Mechanical Damage ........................................126 

Vertebrate Pests ..............................................128 

Pesticide Information ..................................... 130 

Fungicides, Herbicides, Insecticides, and Miticides ...131 

Pesticide Resistance Issues .......................................131 

Pesticide Safety ........................................................131 

Appendixes ..................................................... 133 

A. Pest and Disease Photo Chart ............................134 

B. Biological Controls Photo Chart .........................144 

C. Conifer Species ...................................................148 

D. Seasonal Monitoring Guide ................................150 

E. Insect Trap Use and Construction .......................154 

F. Scouting Record Template for 

Common Pine Pests ............................................160 

G. Scouting Record Template for 

Common Spruce Pests ........................................164 

H. Scouting Record Template for 

Common True Fir and Douglas-Fir Pests .............168 

I. Growing Degree Day and Temperature 

Record Template .................................................172 

J. Pesticide Application Record Template ...............174 

K. Contact Information ...........................................176 

L. Lists of Pests.......................................................182 

—Pests in Alphabetical Order ............................183 

—Pests by Host Genus .......................................184 

Bibliography .................................................... 185 

Glossary ........................................................... 199

INTEGRATED 

PEST MANAGEMENT BASICS 

Christmas tree growers everywhere face obstacles 

to growing healthy trees. Perhaps the largest 

challenge is controlling the pests that feed on 

and live off of conifers. The aim of this manual 

is to help growers combat their pest problems 

by using integrated pest management (IPM) 

methods. IPM is a multilayered approach to 

keeping pests and pest damage to a minimal level 

with the smallest cost to health, environment, 

and budget. 

IPM attempts to improve on some of the 

common problems with traditional Christmas 

tree pest control. One problem is improper timing 

of pesticides. Traditionally, growers may use 

two unsuccessful timing methods: calendar spraying 

and reaction spraying. Calendar spraying is 

making pesticide applications at the same time 

every year regardless of the actual pest situation. 

Reaction spraying is making a spray immediately 

upon discovery of a pest problem. Both of these 

methods are generally incorrect because they 

may not coincide with the target pest’s vulnerable 

life stage. 

Another problem IPM attempts to correct is 

the overuse of broad-spectrum, or nonspecifi c, 

pesticides. Broad-spectrum pesticides kill a wide 

range of insects, not just the target pests. Benefi 

cial insects that help keep pest populations in 

check may be eliminated by these harsh pesticides. 

If spray timing is not correct, growers may 

make multiple broad-spectrum applications and 

possibly still not get control. Using the steps of 

IPM addressed on the following pages, growers 

should be able to implement safer, more effective 

pest control.

STEP 1. Preparing for IPM: 

Planning and Prevention 

IPM practices are generally aimed at an 

established crop, but some pest prevention 

can occur before the crop is even planted. 

Most growers know what species they 

intend to grow at their location. However, 

since a tree grown in a suitable location is 

a good defense against pests, consider the 

following points before planting: 

• Property assessment. Identifying the 

challenges to growing trees on a property 

will help growers determine the 

best locations for tree blocks. Assessing 

the property may include collecting and 

testing soil samples, surveying property 

slope and topography, and determining 

water drainage rates and sunlight levels. 

Historical meteorological information 

and knowledge of past crops can be useful. 

Talking to area growers about possible pest 

problems (insect, mite, disease, mammal, 

etc.) to expect would also be benefi cial. 

• Tree species. Selecting tree species best 

suited to the property is the next step. 

This will involve research and may mean 

that the site is not appropriate for the 

intended species. By using printed publications, 

Web resources, and information 

from other growers and seedling nurseries, 

growers can predict the success of the 

intended crop. For instance, knowing that 

Colorado blue spruce may grow best in a 

fl at, sometimes wet fi eld can help a grower 

decide not to plant Fraser fi r, which could 

be susceptible to root rot issues in this 

same location. 

• Seed source. In addition to choosing 

tree species, growers may have options 

for the seedlings based on seed source. 

Seed sources generally refer to different 

geographical regions where the trees grow 

naturally. The trees resulting from these 

seeds may be adapted to certain growing 

conditions or may show some pest resistance 

in addition to foliage and growth 

attributes. Growers selecting seedlings 

from a particular seed source for its pest 

resistance need to understand that resistance 

characteristics may change when 

the tree is grown outside its natural range. 

• Planting. The success of a tree block has a 

lot to do with proper planting of seedlings 

and transplants. Consider the following 

factors when planting: 

— Spacing. Adequate spacing between 

the trees will provide good air circulation, 

which can prevent disease 

development and allow for good spray 

coverage during pesticide applications. 

Other factors to consider when spacing 

trees are the minimum requirements 

for mowing, harvesting, and shearing 

equipment. A spacing of 5½ feet by 

5½ feet or 6 feet by 6 feet will allow for 

fewer trees than a spacing of 4 feet by 

4 feet, but the trees with wider spacing 

will be healthier through harvest time. 

Spacing needs will vary depending on 

the fi nal desired age of the trees in a 

block. 

— Depth. The depth at which seedlings 

are planted will greatly affect tree 

health. Planting trees so that the root 

collar is at or just slightly below the soil 

line and not planted too deep or too 

shallow will be a factor in preventing 

pest attack. 

— Method. Some planting methods 

are more prone to future problems. 

Mechanical planters may encourage 

J-rooting. In this situation, roots do 

not spread out but actually grow in a 

“J” shape (Fig. 1). As a result, trees are 

unable to establish a good root system, 

which after several years will lead to 

poor tree health. J-rooted trees are also 

more susceptible to insect and disease 

problems. 

Figure 1. J-rooted tree resulting from improper 

planting. Courtesy of Tracey Olson, PDA 

IPM FOR CHRISTMAS TREE PRODUCTION ............................................................................................................................................................................................................................................. 6

STEP 2. Identifi cation and 

Understanding of Pests and 

Problems 

After the Christmas tree block is planted, 

the focus of an IPM approach will be on 

preventing losses due to pest damage. The 

term “pest” may include any organism— 

diseases, insects, mites, nematodes, mammals, 

birds, or weeds—that is detrimental 

to the health of the tree. Environmental 

factors such as air pollution may also cause 

damage. Knowing which pest or what factor 

is causing damage to a tree is essential to 

fi nding a means of stopping or controlling 

that problem. 

Insects and Mites 

The largest groups of Christmas tree pests 

are two classes of arthropod pests, or organisms 

with jointed body parts and an exoskeleton. 

These two classes are the insects 

and the arachnids. Arachnids include mites, 

spiders, and ticks, but only the mites can 

be detrimental to tree health. Experienced 

growers are familiar with the insect and 

mite pests that may affect their trees. They 

have, or know where to fi nd, information 

about the life cycle and habits of these pests 

in order to determine the best method and 

timing for both detection and control, if 

necessary (Fig. 2). 

For example, spider mites (Fig. 3) and 

(eriophyid) rust mites feed openly on tree 

foliage without any extra protection. It is 

easier to control these pests than one that 

spends most of its life covered by a protective 

coating, such as pine needle scale. In 

addition to the mites, pests that feed openly 

include aphids, gypsy moths, and sawfl ies. 

The larger group of insect and mite pests 

includes those that are only exposed for a 

short time during their life cycle. Timing 

sprays to target susceptible life stages is critical 

for effective control. Pests that fall into 

this category include all the scale insects, 

most adelgids, bagworm (Fig. 4), weevils, 

bark beetles, and midges. 

Knowing these and other insects is essential 

for growers to protect their trees from 

pest infestation. In addition, recognizing 

which insects and mites are benefi cial and 

taking steps to avoid killing these organisms 

can limit or even eliminate the need for 

chemical control measures. 

Stinging insects, such as wasps and bees, 

are also a major concern on Christmas tree 

farms, especially during summer mowing 

and fi eld maintenance. These insects are 

frequently attracted to trees infested with 

aphids and scale insects. Recognizing both 

the pest and the stinging insects and understanding 

their biology are important when 

considering employee safety. 

Figure 2. Balsam twig aphid life cycle. Courtesy of Sarah Pickel, PDA (photos courtesy of PDA) 

Figure 3. Spruce spider mite 

feeding openly on tree foliage. 

Courtesy of Rayanne D. Lehman, 

PDA 

Figure 4. Protective casing 

containing bagworms. Courtesy 

of Rayanne D. Lehman, PDA 

IPM BASICS .............................................................................................................................................................................................................................................................................................. 7

Figure 5. Rhabdocline needle 

cast symptoms on Douglas-fi r. 


PDA 

Figure 6. Reddish-brown 

splotches, symptoms of Rhabdocline, 

found in late winter. 

Courtesy of Tracey Olson, PDA 

Figure 7. Buck rub on a young 

fi r. Courtesy of Rayanne D. 

Lehman, PDA 

Diseases (Fungi and Nematodes) 

Numerous diseases of Christmas trees are 

caused by fungi and nematodes, with the 

majority being caused by fungi. Fungi are 

simple organisms that survive in soil, water, 

and dead or living organic material. Cool 

temperatures, high moisture, and low air 

circulation will encourage fungal development. 

In Christmas tree plantations, these 

factors often occur together, making disease 

development common. Symptoms include 

swelling or shrinking of the wood tissue, 

discolored or deformed needles, early needle 

drop, and wilted foliage. The disease types 

are referred to as cankers, galls, needle casts, 

rusts, rots, and blights. 

Fungal diseases spread from tree to tree 

by microscopic spores. At a specifi c stage in 

disease development, mature fruiting bodies 

on the diseased host release the spores. 

Wind, rain, or other physical contact will 

spread these spores to other trees, initiating 

a new disease cycle. Growers should be 

aware of the diseases that may affect their 

trees and have an understanding of the 

symptoms (Figs. 5 and 6). 

Unlike the insect and mite pests, 

diseases are not usually controlled—they 

are prevented. Fungicides will not kill the 

fungus but only protect the plant tissue from 

infection. Therefore, most fungicides must 

be applied before spores would contact the 

healthy tissue. 

Less common are the diseases caused by 

nematodes. Nematodes are microscopic, 

simple worms. They damage plants by 

sucking out the contents of plant cells. In 

the case of pine wilt disease, a wood-boring 

beetle transfers nematodes to the tree. 

Vertebrates (Mammals and Birds) 

Vertebrates are animals with backbones. 

The vertebrates of concern for Christmas 

trees are birds and mammals, especially deer 

and rodents. Birds may damage tree leaders 

by perching on the tender growth, resulting 

in broken or bent leaders. Deer may severely 

damage shoot tips by feeding on them. 

Bucks can also cause injury by rubbing their 

antlers on the main trunk (Fig. 7). Rodents 

may kill trees by girdling the lower trunk as 

they feed on the bark. 

Weeds 

Weeds are undesirable plants that interfere 

with the growth of a crop such as Christmas 

trees. Generally herbaceous, they reproduce 

and spread through seeds and rhizomes 

(underground stems). Weeds compete with 

trees for soil nutrients, water, and sunlight 

(Fig. 8). They inhibit good air circulation 

between trees—a contributing factor for 

disease development. Weeds also hinder 

the movement of people and equipment 

through and between rows, making good 

pesticide coverage diffi cult. Through good 

mowing practices and the assistance of 

herbicides, growers can control weeds to a 

manageable level. 

Weed control will not be covered in this 

manual. Information on weeds can be found 

in the Penn State publication Controlling 

Weeds in Nursery and Landscape Plantings 

by Dr. Larry Kuhns. Growers can also look 

to their state land-grant universities for additional 

weed resources. 

Environmental Factors 

Injury to Christmas trees is not solely a result 

of living organisms. Environmental factors 

can also have a negative effect. Damage 

from severe weather can be equally as bad 

or worse than a disease or insect problem. 

Extreme cold, wind, or hail (Fig. 9) can 

cause needle discoloration, shoot dieback, or 

lesions on stems and twigs. Some unnatural 

environmental factors also exist. Pesticides 

and fertilizers applied at the wrong time of 

year or in the wrong amounts will also damage 

the trees. Air pollution, although hard to 

identify, may also harm trees. 

Figure 8. Common milkweed, a familiar nursery 

weed in the Mid-Atlantic and Northeast United 

States. Courtesy of Theodore Webster, USDA 

Agricultural Research Service, Bugwood.org 

(#1552251) 

Figure 9. Hail damage 

on eastern white 

pine. Courtesy of 

Rayanne D. Lehman, 

PDA 

IPM FOR CHRISTMAS TREE PRODUCTION ............................................................................................................................................................................................................................................. 8

STEP 3. Monitoring Trees for Pest 

Populations 

Monitoring is the key to any successful IPM 

plan. On farms not using IPM, the process 

of pest control usually involves pesticide 

applications based on a calendar date. These 

types of applications are made regardless of 

the verifi ed presence of the target pest. Another 

type of pesticide application on farms 

not using IPM is the “see and spray” method: 

spray only when the pest or damage is seen. 

At times, applications are made based only 

on damage, not on actual pest presence. IPM 

looks at pest management differently. Monitoring 

to determine whether there is a need 

to take action is more effective than either 

of the methods above. 

Monitoring provides growers with an 

accurate picture of pest situations on their 

Christmas tree farms. Monitoring includes 

the use of traps that catch pests to indicate 

their presence as well as direct observation 

by a scout. The process involves regular, 

close inspection of the trees on the farm for 

symptoms or signs of pest activity (Fig. 10). 

“Symptoms” refer to the damage or evidence 

of activity, such as yellowed needles 

or wilted shoots; “signs” refer to the actual 

organism causing the damage, such as black 

fruiting bodies on the underside of a needle 

or a bagworm case hanging from a branch. 

By inspecting for these symptoms and signs, 

noting the stage of development, and 

evaluating the level of infestation or infection, 

growers can create an individualized, 

accurate, and timely plan of action for each 

fi eld. Monitoring can be done by a farm employee 

whose time is committed to scouting 

activity or by a professional scout hired as a 

consultant. The key is to scout regularly, not 

just once or twice a season. 

Figure 10. Scout conducting his 

weekly activities. Courtesy of 

Cathy Thomas, PDA 

Growing Degree Days/Temperature 

One method of monitoring specifi c pest populations 

is growing degree day (GDD) tracking. 

For insects, mites, and plants, development is 

infl uenced by daily heat accumulations from 

ambient air. (Disease progression is based 

on other factors such as host plant development, 

humidity, leaf wetting, etc.; therefore, 

the GDD method does not apply.) Tracking 

heat accumulation by monitoring daily air 

temperatures has proved useful in predicting 

the appearance of certain pest life stages. 

The growing degree day method uses the 

daily average temperature (using low and 

high temperatures for 24 hours) to determine 

the heat unit exposure of the pest or 

plant. The GDDs accumulate daily until the 

required amount of heat for an event (hatch, 

emergence, pupation, etc.) is reached. This 

is not a specifi c number, but a range. For 

instance, spruce spider mite overwintering 

eggs begin to hatch around 50–121 GDDs. 

A scout can effectively monitor for a pest by 

knowing the range of GDDs for the target 

stage of each pest. Known GDD ranges for 

many Christmas tree pests can be found in 

the Growing Degree Days Chart on page 18 

and on each individual pest fact sheet. 

As a general rule, GDD recording should 

begin on March 1. Although each organism 

has a unique base temperature, most GDD 

calculations use a base temperature of 50°F. 

The base temperature is the minimum temperature 

required by an organism for development 

to proceed. Temperature should be 

tracked using a minimum (low)/maximum 

(high) thermometer capable of storing readings 

for several days. After calculating each 

day’s GDD value, add it to the accumulated 

total. Never include negative numbers. 

GDD FORMULA 

Low Temperature + High Temperature – 50°F = Daily GDD 

2 

(Average temperature (base 

for a 24-hour period) temp.) 

IPM BASICS .............................................................................................................................................................................................................................................................................................. 9

Figure 11. A minimum/maximum 

thermometer collecting 

temperature data in a fi eld. 

Courtesy of Cathy Thomas, PDA 

Figure 12. A probe thermometer 

measuring soil temperature 

at a depth of approximately 

2 inches. Courtesy of Cathy 

Thomas, PDA 

DAILY WEATHER LOG 

Air Temperature Daily 

GDD 

Running 

Date Time Low High Ave. GDD Total 

3/24 5:30 pm 45°F 73°F 59°F 9 9 

3/25 5:30 pm 35°F 55°F 45°F 0 9 

3/26 5:30 pm 46°F 60°F 53°F 3 12 

March 24 

45 + 73 = 118 = 59 – 50 = 9 GDD Count positive number. 

2 2 

March 25 

35 + 55 = 90 = 45 – 50 = -5 GDD Do not add negative 

2 2 numbers. 

March 26 

46 + 60 = 106 = 53 – 50 = 3 GDD Add positive number 

2 2 to total. 

Remember that temperatures may differ 

in different blocks. A tree block on a southfacing 

slope will experience higher temperatures 

than a block on a north-facing slope, 

which can result in several day differences 

in pest development. 

Using minimum/maximum thermometers 

(Fig. 11) is the least expensive method of 

gathering data to calculate GDDs. Several 

other types of equipment will gather GDD in 

the fi eld. These are commonly called biophenometers, 

data loggers, or weather monitors. 

The prices vary, but they are generally more 

expensive than a simple minimum/maximum 

thermometer. Weather services, both paid 

and free, can also provide required data. 

Skybit is a subscriber-based service located 

in Pennsylvania (see www.skybit.com). The 

National Weather Service (NOAA) provides 

daily low and high temperatures from 

weather stations. NOAA data will not be 

specifi c to your area and should not replace 

on-farm temperature collection. 

Soil Temperature 

Research conducted in Pennsylvania on 

white pine weevil (Pissodes strobi) has 

shown that collection of soil temperatures 

is equally or more effective than GDD in 

determining the weevil’s spring emergence 

time. Temperatures were taken with a probe 

thermometer from soil beneath a tree in 

the target block. The probe was inserted 

to a depth of 2 inches on the sunny side of 

the tree (Fig. 12). Once the soil temperature 

reached 50°F, weevils could be found 

in traps and on tree leaders. This method 

may also be effective on other insects that 

overwinter in soil. 

Personal Monitoring Kit 

When monitoring, a scout must have a few 

very important tools. Keep these tools together 

in a backpack or bag for convenient 

transport to the fi eld on each visit (Fig. 13). 

HAND LENS 

This small magnifi er allows a scout to see insects, 

mites, and fungal fruiting bodies that 

would otherwise be too small to see (Fig. 

14). These can be purchased in magnifi cation 

strengths of 10X, 15X, and 20X. The 

15X hand lens is a good strength to use and 

still has a reasonably sized fi eld of view (see 

Appendix K for supplier information). 

Figure 13. Scouting tools and convenient zipper 

pack for storage. Courtesy of Brian Schildt, PDA 

Figure 14. The number one 

tool for a scout: a hand lens. 


NOTEBOOK AND PEN 

Use these to record symptoms, signs, 

population stage and level, damage severity, 

fi eld or weather conditions, and location. 

This information can be a reference for the 

current season as well as a resource that 

shows pest activity from previous seasons. 

Suggested forms for scouting observations 

and notes are included in the Appendix 

section of this manual. 

FLAGGING TAPE 

Use brightly colored fl agging tape to mark 

symptomatic trees so they can be readily 

located at a later date. Use a permanent 

marker to record specifi c information on the 

tape. Remember to use a color other than 

those used for digging or cutting purposes. 

IPM FOR CHRISTMAS TREE PRODUCTION ........................................................................................................................................................................................................................................... 10

CLIPBOARD/FLAT SURFACE 

Hold a clipboard or another fl at surface 

under a branch to collect mites, aphids, 

or other insects dislodged by tapping the 

branch (Fig. 15). The back of a clipboard 

that has been painted to have one half 

black and one half white is very useful when 

beating trees to detect pests. The inside 

back cover of this manual is a good fl at 

surface to use. 

PRUNERS/POCKET KNIFE 

Growers may need to clip symptomatic 

twigs or branches to observe them more 

closely for signs of insects, mites, or disease 

spores. Collecting samples for identifi cation 

by a diagnostic expert (Fig. 16) may also be 

necessary. 

BOTTLES/PLASTIC BAGS 

Occasionally, growers need to collect samples 

to examine at a later date or send to a 

diagnostic laboratory. In addition to bottles 

and plastic bags for collecting samples, 

having a method of labeling the specimens 

(permanent marker, adhesive labels, etc.) is 

also helpful. Any sample collected in alcohol 

should be labeled with a pencil-written 

tag inserted into the vial. 

Monitoring Tips 

1. During the growing season, a scout 

should be monitoring fi elds weekly to 

look for evidence of pests, diseases, and 

other problems. 

2. If possible, scout on cloudy days. The 

muted sunlight makes it easier to observe 

chlorotic, or yellowed, symptoms 

in the fi eld. Some pests are also more 

visible in subdued light. 

3. When monitoring blocks for problems, 

scouts should walk through the fi eld 

while keeping an eye out for obvious 

problems. They should also select nonsymptomatic 

trees in a random pattern 

to inspect more closely. For these trees, 

be sure to look at the inside for discoloration. 

4. Prune a few twigs from the interior 

and lower portion of the tree for closer 

inspection. 

5. Tapping branches over a fl at surface 

will dislodge insects and mites, making 

them easier to see (use the tapping 

sheet at the back of this manual). 

Trapping 

Several types of insects can be monitored 

using traps. The traps are usually baited with 

pheromones (insect-produced attractants) 

or other chemicals (Fig. 17) and placed in 

the fi eld. They are designed to hold the 

insect until the trap can be checked by 

a scout. When monitored regularly, the 

collection data can help with the timing 

of control applications. Traps are used for 

monitoring purposes, not control. For more 

information on trapping, see Appendix E. 

Sticky cards are another type of monitoring 

tool used to trap insects. These 

adhesive-coated index cards, which come 

in various sizes and colors, may be used 

to monitor the emergence of small pests 

(Fig. 18). For example, in the case of some 

scale insect pests, 3-by-5-inch sticky cards 

are used to attract the adult male life stage. 

The cards are attached with clothespins 

to scale-infected tree limbs before the 

expected time of emergence. Cards can 

then be monitored regularly. 

Figure 18. Yellow sticky cards used to trap 

fl ying adult male scales. Courtesy of Cathy 

Thomas, PDA 

Diagnosis 

Proper identifi cation of the pests that 

have been found through the monitoring 

process is an important next step in the 

pest management process. With training, 

experience, and the use of printed and Web 

resources and guides, growers or scouts can 

usually make an accurate diagnosis. For 

diffi cult samples, consult an expert or a 

diagnostic lab. Regional inspectors for the 

state department of agriculture and county 

extension educators can be very helpful in 

obtaining necessary expert assistance. For 

contact information for select agencies, see 

Appendix K. 

Figure 15. Tapping branches 

over a white notebook or fl at 

surface to dislodge and observe 

pests. Courtesy of Brian Schildt, 

PDA 

Figure 16. A good set of pruners. 

Courtesy of Joseph O’Brien, 

USDA Forest Service, Bugwood 

.org (#5044044) 

Figure 17. White pine weevil 

trap. Courtesy of Cathy Thomas, 

PDA 

IPM BASICS ............................................................................................................................................................................................................................................................................................ 11

STEP 4. Setting an Action 

Threshold 

The action or control threshold refers to the 

population level for a specifi c pest at which 

some control measure is justifi ed in order 

to avoid economic loss or aesthetic damage 

to a crop. Christmas trees can sustain some 

damage and still be aesthetically acceptable 

to consumers. However, when the damage 

becomes unacceptable, the crop has passed 

what is known as the aesthetic injury level. 

The action threshold will be exceeded 

before the aesthetic injury level has been 

reached. 

Because pests vary in the severity of 

damage they cause, action thresholds will 

also vary. A pest that has great potential 

to damage or disfi gure a tree, such as white 

pine weevil, will have a lower threshold 

than a more minor pest, such as the balsam 

twig aphid, for which some damage may be 

acceptable. Action thresholds are further 

discussed in the individual fact sheets found 

in this manual. 

The question of acceptable action 

thresholds will also vary depending on how 

the trees are marketed. If trees are sold as 

nursery stock, often referred to as balledand-burlapped 

(B&B) trees for the way 

they are dug and bound, then the issue of 

threshold is no longer solely determined by 

the grower. All nursery stock in Pennsylvania 

is subject to state regulation, and the 

level of pest tolerance for the trees is based 

on a regulatory threshold. These regulatory 

thresholds are enforced by state or federal 

nursery inspectors. Some of these thresholds, 

such as those for pests under state or 

federal quarantines, are zero tolerance. Cut 

Christmas trees in Pennsylvania are not subject 

to regulation. However, if trees are to be 

shipped out of state, federal quarantines will 

be enforced. 

Growers need to be aware of these 

regulatory issues when considering control 

actions on their farms. More information on 

state regulatory requirements can be found 

in the Conifer Certifi cation and Quarantine 

Guidelines section on page 16. 

STEP 5. Selecting Control Options 

IPM does not usually rely on only one 

method or tactic. Often, growers employ a 

combination of methods to achieve a wellrounded, 

long-lasting management program. 

Cultural and Mechanical Control 

Best growing practices and physical methods 

of control are often the fi rst defense to 

pest issues and can be the least expensive. 

Examples of these controls include: 

• Mowing. Through consistent management 

of grasses and weeds by mowing, 

growers are able to facilitate good air 

circulation through tree blocks. Good air 

circulation helps prevent disease development 

and opens the lower portion of the 

trees to receive chemical applications, if 

necessary. 

• Pruning. In addition to shaping trees, 

pruning can also help eliminate some 

pest issues. Pruning is one way to help 

eliminate the galls produced by adelgids 

or needles infested with needle midges. 

Growers may eliminate some of the 

feeding damage produced by balsam twig 

aphids, bagworm, or pine sawfl ies. Also, 

clipping out leaders infected by white 

pine weevil is an effective management 

method. Butt-pruning, or the removal of 

the lowest whorl of branches, opens the 

bottom of the tree to provide better air 

circulation, weed control, and pesticide 

application. It also permits heating of the 

soil and may make the area less attractive 

to certain insect and rodent pests. 

• Culling. This is the practice of removing 

weakened, infected, and dying trees from 

the fi elds. Eliminating these reservoirs 

from the fi elds not only removes a source 

of infestation but will also allow for better 

air circulation among the remaining trees. 

It is an effective, low-cost method for 

reducing a recently introduced pest that 

has the potential to cause serious damage. 


Biological Control 

Biological control, or biocontrol, is the use 

of living organisms (parasitoids, pathogens, 

predators) to control diseases, insect pests, 

and weeds (Fig. 19). Biocontrol plays an 

important role in IPM but does not defi ne 

it. Many IPM techniques are geared toward 

enhancing or preserving biocontrols. A 

successful IPM program combines many 

methods aimed at keeping pest populations 

below damaging levels. 

In Christmas trees, no biocontrol agents 

exist for disease, and those for weed control 

are poorly understood. But numerous 

naturally occurring parasitoids, pathogens, 

and predators of insects and mites are present 

(Fig. 20). Growers are often not aware 

of these natural enemies and continue to 

rely on insecticides to control pests. Excessive 

pesticide use may eliminate natural 

enemies, further increasing reliance on 

pesticides. The goal of a good IPM program 

is to recognize and preserve these biocontrol 

agents. The benefi t is often a reduction of 

pesticides and costs. 

In order for natural enemies to survive, 

they must not be 100 percent effi cient. A 

small number of pests are always needed to 

keep the natural enemies around. Because 

the pests are not eliminated, biocontrol is 

not as popular when dealing with a crop 

such as Christmas trees where aesthetics 

can be important. However, biocontrol 

agents are extremely useful if dealing with 

a pest that either has a high threshold of 

damage or can be tolerated in a low-level 

population for a long time. 

Not all pests are suitable for biocontrol. 

Borers, for instance, must be treated as soon 

as they are detected. White pine weevil 

has very few natural enemies, and none are 

known for pine root collar weevil. Some 

pests, such as aphids and mites, are easier to 

keep in check with natural enemies, but this 

is only applicable if regular scouting is used to 

monitor both the pest and benefi cial populations. 

By monitoring, growers can take action 

if the pest populations are not being kept in 

check by their natural enemies. 

Some research has been done with 

insect-pathogenic nematodes that grow in 

the bodies of their hosts (e.g., beetle larvae), 

eventually killing them. Neoaplectena 

carpocapsae (Weiser) has been extensively 

studied and tested, and Heterorhabditis heliothidis 

is receiving attention. In order for the 

nematode to do its job, it must occur with a 

bacterium since neither can survive alone. 

Sometimes the nematode-bacterium is not as 

effective as an insecticide, but that depends 

on the situation. The most critical factor 

restricting wide use of insect-pathogenic 

nematodes is overcoming nematode environmental 

sensitivity. Nematode success is 

greatest when moisture and temperature 

are at the optimum level for that location. 

Active research continues at universities 

and in private industries to increase the effectiveness 

of insect-pathogenic nematodes 

and make them a more cost-effective tool. 

Chemical Control 

When biocontrols are not suitable, not 

working as they should, or overwhelmed by 

a pest outbreak, chemical control is the next 

step. “Soft” or “reduced-risk” pesticides, 

such as insecticidal soap, ultra-fi ne horticultural 

oil, and neem compounds, are used 

for treating outbreaks that happen while 

managing with biocontrols. Consult with 

the biocontrol distributor before spraying to 

ensure compatibility. 

The best control plan treats the pest 

populations with minimal disruption to 

the natural enemies. This can be accomplished 

several ways, including alternating 

insecticide classes, altering spray timing, or 

adjusting cultural practices. Another approach 

is to use soft insecticides that are not 

harmful to benefi cial insects. Horticultural 

oils are effective in killing overwintering 

eggs and sessile insects but do little harm to 

natural enemies. Caterpillars, but not sawfl y 

larvae, can be controlled with the bacterial 

insecticide Bacillus thuringiensis, or Bt. Some 

pest-specifi c insect growth regulators (IGRs) 

are also effective. There are three general 

types of chemical control: biorational, 

horticultural oil, and traditional. 

BIORATIONALS 

As pesticide regulations increase and the 

general public gains awareness of pesticides 

used on Christmas tree farms, more attention 

is directed toward alternatives. Alternatives, 

otherwise known as biorational 

products, include insecticidal soap, pyrethrum, 

and rotenone (not an oil). 

Insecticidal Soap 

The insecticidal action results when fatty 

acids from plant oils are combined with the 

potassium bases to produce potassium salts 

of fatty acids. The inactive ingredients are 

water and alcohol. 

Figure 19. A praying mantis, a 

generalist predator. Courtesy of 


Figure 20. Multicolored Asian 

lady beetle larva, an aphid 

predator. Courtesy of Cathy 

Thomas, PDA 

IPM BASICS ............................................................................................................................................................................................................................................................................................ 13

Advantages: 

1. The mode of action (MOA) of insecticidal 

soap is physical, so pests are less 

likely to develop resistance. Soap acts 

on contact, penetrating body and cell 

membranes and causing quick death. 

2. Insecticidal soap has very few long-term 

negative effects on benefi cial insects. 

3. Insecticidal soap is relatively nontoxic 

to wildlife and humans. 

4. It can be applied using a variety of 

application equipment, and disposal of 

unused material and containers is easy. 

Disadvantages: 

1. The main drawback of using insecticidal 

soap is the lack of residual activity; 

therefore, more frequent applications 

are required. 

2. The effectiveness of insecticidal soap 

decreases when mixed with hard water. 

3. Soap will kill natural enemies on 

contact when plant surfaces are wet. 

Neem Seed Extracts 

Neem seed extracts are botanical or plantderived 

pesticides. Neem oil extracts are 

from seeds of the neem or margosa tree 

(native to Asia). Azadirachtin, one of the 

active ingredients in neem oil, has growthregulating 

properties. Neem oil has more 

than 12 other active ingredients that act 

as egg-laying deterrents, feeding inhibitors, 

growth regulators, repellents, sterilizing 

agents, or toxins. 

Advantage: 

1. Various neem products affect more than 

170 species of insects, mites, and nematodes 

that infest households, humans, 

livestock, plants, and stored products. 


1. The raw neem oil has an unpleasant 

garlic-sulfur smell, which is often 

eliminated in commercial production. 

2. Natural enemies of plant pests are 

slightly affected by these products. 

3. Effectiveness may be questionable. 

Bacillus thuringiensis (Bt) 

Bt is a bacterial or microbial insecticide 

that attacks the gut of Lepidoptera larvae. 

The caterpillars ingest the Bt spores, which, 

in the alkaline conditions of the insect’s 

gut, release toxins. This causes the insect’s 

own digestive juices to attack the gut wall, 

creating holes that allow the contents to 

poison the body. Death may take a few days, 

but the insect stops feeding soon after the 

Bt is ingested. It is best applied when the 

caterpillars are young. Other formulations 

of Bt exist that target other insect groups, 

but none have been successfully used in 

Christmas tree production. Bt is used to 

control caterpillars and has no effect on 

sawfl y larvae. 

Bacillus popilliae 

Better known as milky spore disease, Bacillus 

popilliae was the fi rst commercially introduced 

microbial insecticide. It has been 

used for many years to control Japanese 

beetle larvae in soil. The spores will remain 

dormant in the soil until future generations 

of Japanese beetle larvae are present. It is 

not effective against other white grubs. 

HORTICULTURAL OILS 

Horticultural oils work in two ways. The 

fi rst MOA is suffocation. When the oil coats 

the insect’s body, it prevents the exchange 

of gases in the pest. Eggs are also affected in 

this manner. The second MOA is penetration 

of the insect’s external membranes and 

subsequent entrance into the individual 

cells, where cell functions are disrupted. 

Both dormant oil and the highly refi ned 

oils for use during the growing season are 

available. 

Advantages: 

1. Economical—most oils are less expensive 

than pesticide products. 

2. There are no proven cases of insects or 

mites becoming resistant to oil. Even 

insecticide-resistant species are susceptible 

to oil. 

3. Oils are environmentally and applicator 

friendly, posing little hazard with their 

use. 

4. When used to control overwintering 

pests, growers have more opportunities 

to select a day to make an application. 

5. Early season feeding damage of overwintering 

pests will be eliminated or 

signifi cantly reduced. 

6. A wide assortment of pests can be 

controlled on a large number of hosts. 

7. Oil has no residual effect and will only 

harm the organism by direct contact. 



1. Oils must come in contact with the 

targeted pest to be effective. 

2. Field conditions and weather during 

applications may limit effi cacy. 

3. Eggs will only be controlled if they are 

present before the oil application. 

4. Oil has no residual effect. 

Cautions: 

1. When temperatures are below freezing, 

oil can cause phototoxicity. Under 

these conditions, the oil and water cannot 

stay mixed and the water freezes, 

allowing oil droplets to accumulate. 

When thawing does occur, the water 

will evaporate, concentrating the oil on 

plant surfaces. 

2. Oil can also cause phototoxicity when 

temperatures are above 90°F. Burning is 

also a concern on the sunny side of the 

trees or during times of drought. 

3. Apply oils when conditions allow for 

prompt drying; be sure to avoid drift or 

overspray. 

4. Mistaken dormancy is a problem in 

early fall or late spring. In early fall, if 

the plant is not in dormancy and the 

leaves are water defi cient, then the oil 

will burn the foliage. In late spring, new 

growth is burned if the dormant period 

ends before the oil is applied. 

5. Conifers known to be sensitive to 

dormant oils include Douglas-fi r and 

spruce. Be sure not to spray oil on 

glaucous, or blue, varieties of conifers 

because the blue color will be removed 

and may not return for 2–3 years. 

Genetic variability may affect individual 

plants differently, even if the variety 

is known to be tolerant to oil. 

When using oil, as with any pesticide, 

be sure to follow label directions. Some 

oils can be mixed with other pesticides to 

increase the level of toxicity. Never mix 

oils with dimethoate or any type of sulfur, 

as serious phytotoxic reactions will result. 

Most fungicides are not compatible with 

oils. Always consult the oil and insecticide 

labels for compatibility. Always choose 

“superior-type” oils to provide further 

insurance that these products are safe. 

TRADITIONAL 

This largest group of pesticides includes 

insecticides, miticides, fungicides, and 

herbicides. They can be the most effective 

materials available to prevent and destroy 

pests; however, they are frequently used 

when other options are available. For many 

years, broad-spectrum insecticides were the 

only tools available to growers. In recent 

years, newer, soft materials targeting specifi c 

groups of pests have been introduced. These 

materials are less likely to disrupt natural 

enemies while keeping the target pest in 

check. 

The use of traditional pesticides should 

always be a last resort. If a traditional 

pesticide application is a must, choose the 

right product, time, and dosage to avoid 

problems. Inappropriate use may rid the 

area of pests now, but it may also cause a 

more serious problem later as pests become 

resistant and natural enemies are depleted. 

For maximum benefi t and to avoid hazards, 

choose formulations that pose the least 

threat to nontarget species. Be sure to rotate 

classes of pesticides to decrease the chances 

of pests developing resistance. Always read 

the label before using any pesticide product. 

IPM BASICS ............................................................................................................................................................................................................................................................................................ 15

Step 6. Evaluating Results of 

Control Actions 

The ultimate goal of any IPM program is 

to keep pests under control. Not all control 

measures will be met with success. Sudden 

rain showers, rapid drop in temperature, 

inadequate coverage, and poor mixing are 

only a few of the causes of control failures. 

In addition, pest populations may rebound 

quickly despite the best techniques of an 

applicator. Monitoring pests after a control 

application is equally as important as 

monitoring them before the application. 

Scouts should check the target pests 3–5 

days after any application. Some materials 

may require a longer time period to be effective. 

Checking effectiveness of fungicides is 

very diffi cult, but trees should be examined 

to make sure a spray residue is on all areas of 

the tree. Heed reentry interval requirements 

on the pesticide label when scouting after 

an application. 

If the target pest is still present in suffi 

cient numbers to cause damage, a repeat 

application may be necessary. Consider the 

following factors when determining the 

need for reapplication of pesticides: 

• Pest life cycle: 

— Vulnerability of certain life stages 

— Timing of generations 

— Number of generations per season 

• Pesticide properties: 

— Mode of action 

— Residual effect 

• Weather conditions following the 

application 

• Number of applications permitted 

according to the pesticide label 

Even the best method of control will not 

eliminate the need for continued monitoring 

of pests. Many pests are capable of 

reappearing at a later date. They may be 

blown in from neighboring fi elds or woodlots, 

arrive with new seedlings, or have been 

present in an undetectable level during 

regular scouting. 

Pennsylvania Conifer Certifi cation 

and Quarantine Guidelines 

Note: Growers in other states should consult 

with their regulatory authority for their state 

quarantine guidelines. 

Conifer/Christmas tree growers who plan 

to sell B&B trees, seedlings, and transplants 

must have a Pennsylvania state nursery 

certifi cate. Pennsylvania’s Plant Pest Act 

requires that all nursery stock (i.e., plants 

with roots) must be inspected and certifi ed 

as apparently free of injurious plant pests 

before being removed from the growing site. 

To obtain a nursery certifi cate, contact the 


Bureau of Plant Industry Regional Offi ce 

nearest you to request an application. 

The inspector assigned to your county will 

inspect your trees, give recommendations 

for controls of any pest or disease problems 

found, and certify trees suitable for digging 

as nursery stock. This applies to retail or 

wholesale, including local sales. 

Any conifer grower who ships trees 

across state lines should be aware of federal 

and state quarantines that may apply to cut 

and B&B trees, seedlings, and transplants. 

Federal quarantines include the gypsy 

moth quarantine. Under this quarantine, 

all cut and B&B trees of any species moving 

from quarantined areas into or through 

nonquarantined areas must be inspected and 

certifi ed free of all gypsy moth life stages. 

Movement of trees within the quarantined 

area does not require gypsy moth certifi cation. 

Maps of the quarantined areas may 

be found on the USDA Web site at www 

.aphis.usda.gov. 

Pine shoot beetle is another federal 

quarantine that affects the movement of all 

Pinus species—eastern white, Scotch, Austrian, 

and so on. All cut and B&B pine trees 

or any parts, such as branches, wreaths, etc., 

moving from a quarantined to a nonquarantined 

area must be inspected and certifi ed 

free of pine shoot beetle. Maps of the pine 

shoot beetle quarantine zone may also be 

found at www.aphis.usda.gov. 


Each individual state of the United 

States also has its own list of quarantine 

pests. Some examples include the following: 

1. Cereal leaf beetle in California—movement 

of Scotch, red, and Austrian pine 

trees from Pennsylvania to California is 

regulated. 

2. European pine shoot moth in California, 

Hawaii, Idaho, Montana, Nevada, 

and Oregon is regulated. 

3. Hemlock woolly adelgid in Maine, 

Michigan, New Hampshire, Ohio, Vermont, 

and Wisconsin—some of these 

states do not allow any hemlock to be 

shipped there, while others will allow 

them if they meet guidelines. 

4. Japanese beetle—there are four categories 

of states based on whether they are 

considered infested or noninfested: 

• Category 1, Uninfested/Quarantine 

Pest: Japanese beetle is not known to 

occur and is considered a high/moderate-risk 

pest. These states consist 

mainly of the western United States 

and require quarantine certifi cation 

in order to ship nursery stock to them 

from Pennsylvania. 

• Category 2, Uninfested or Partially 

Infested/Regulated Nonquarantine 

Pest: Japanese beetle is not known 

to occur but is established in limited 

areas of the state and is likely to 

spread. The impact can be mitigated 

to an acceptable level by applying 

a treatment before shipping. These 

states consist mainly of the midwestern 

United States. 

• Category 3, Generally Infested and 

Partially Infested: This includes 

states where infestations are generally 

widespread. Since Pennsylvania is 

considered Category 3 (infested), regulations 

for midwestern and western 

states require that a special inspection 

be performed or a chemical treatment 

option for Japanese beetle grubs be 

performed before B&B trees, transplants, 

or seedlings may be shipped 

there. The USDA has enacted a 

Japanese Beetle Harmonization Plan 

designed to slow the artifi cial movement 

of Japanese beetles from the 

eastern United States to the midwestern 

and western states. This plan 

specifi es each state’s requirements for 

shipments, which must be documented 

by a plant inspector before 

shipment may be sent. Generally, the 

East Coast states are in this category. 

• Category 4, Historically Not Known 

to Be Infested/No Regulatory Signifi - 

cance: Japanese beetle spread is not 

likely or the insect is not likely to 

survive or become a pest. The states 

of Florida and Colorado fall into this 

category. 

5. Bagworm in Delaware—the state of 

Delaware includes cut Christmas trees 

in their defi nition of nursery stock. Any 

cut or B&B trees must be inspected and 

found free of bagworm and gypsy moth 

in particular before any shipments are 

made to Delaware. 

The regulations for federal quarantines 

and each individual state’s quarantines may 

be found on the Web at nationalplantboard.org. 

State inspectors and the plant 

protection specialist from the Pennsylvania 

Department of Agriculture can provide 

information about these quarantines. 

IPM BASICS ............................................................................................................................................................................................................................................................................................ 17

This chart provides a list of growing degree 

day ranges (see page 9 for explanation) that 

correspond with a particular pest life stage 

or life cycle event that is critical in the control 

of that pest. This information combined 

with scouting observations will help growers 

achieve the best control of pest problems on 

their farms. 

Growing Degree Day Ranges for Christmas Tree Pests 

Pest Name Degree Day Range Life Stage Comments 

Bagworm 650–750* Larvae emerge from bags 

Balsam twig aphid 30–100 Overwintering eggs hatch 

Conifer rust mites (eriophyid) 7–22 Overwintering eggs hatch 

Cooley spruce gall adelgid 22–81 Spring control of overwintering stage 

2,800–3,000 Fall control of overwintering stage 

Cryptomeria scale 600–800* First-generation crawlers emerge 

1,750–2,130* Second-generation crawlers emerge 

Douglas-fi r needle midge 200–400* Adults emerge from soil 

Eastern pine weevil 7–100 Overwintering adults become active 

Eastern spruce gall adelgid 22–170 Spring control of overwintering stage 

2,800–3,000 Fall control of overwintering stage 

Elongate hemlock scale 360–700 Crawlers fi rst become active 

European pine sawfl y 78–220 Overwintering eggs hatch; larvae present 

Gypsy moth 90–448 Overwintering eggs hatch; larvae present 

Introduced pine sawfl y 400–600 Overwintering eggs hatch; larvae present 

Pales weevil 7–121 Overwintering adults become active; 

treatment to prevent egg laying 

Pine bark adelgid 22–58 Spring control of overwintering stage 

Pine needle scale 298–448 First-generation crawlers emerge 

1,290–1,917 Second-generation crawlers emerge 

Pine root collar weevil 300–350 Overwintering adults become active 

Pine shoot beetle 450–550 New adults emerge 

Redheaded pine sawfl y 400–600 Overwintering eggs hatch; larvae present 

Spruce spider mite 50–121* Overwintering eggs hatch 

Striped pine scale 400–500 Eggs hatch and fi rst crawlers emerge 

(Toumeyella sp.) 

White pine weevil 7–58 Overwintering adults become active 

Zimmerman pine moth 121–246 Larvae emerge from overwintering sites 

*Based on observations in Pennsylvania. 

Monitoring of GDDs begins March 1. 


This calendar gives a range of weeks of 

occurrence for each pest and pest event. 

The purpose of this calendar is to show 

growers how dates of particular pest events 

relate to others throughout the growing 

season. Because only weeks are given rather 

than specifi c dates, these are just approximate 

ranges. The information in this calendar 

was collected from 10 years of scouting 

Insect and Mite Calendar 

Pest Event 

Bagworm larvae emerge 

Balsam twig aphid eggs hatch 

Cooley spruce gall adelgid waxing over 

Cryptomeria scale crawlers emerge 

Douglas-fi r needle midge adults emerge 

Eastern spruce gall 

adelgid 

waxing over 

Eastern pine weevil adults emerge 

Elongate hemlock scale crawlers emerge 

Eriophyid mite* eggs hatch 

Gypsy moth larvae emerge 

Pales weevil adults emerge 

Pine bark adelgid crawlers emerge 

Pine needle scale crawlers emerge 

Sawfl ies larvae emerge 

Spruce spider mite* eggs hatch 

Striped pine scale crawlers emerge 

White pine weevil adults emerge 

*Population resurgence in September may require treatment, if numbers are high. 

early/late emergence 

peak emergence 

data in south-central and southeastern 

Pennsylvania, so these ranges will differ 

from the actual calendar dates in other areas 

of the country. This information should be 

combined with growing degree day calculations 

and scouting data to determine local 

occurrence. Bud break occurs from April to 

May and signals emergence of various insects, 

as well as infection periods of disease. 

Growing Season (weeks) 

March April May June July August 

1 2 3 4 1 2 3 4 1 2 3 4 1 2 3 4 1 2 3 4 1 2 3 4 

IPM BASICS ............................................................................................................................................................................................................................................................................................ 19

PEST FACT SHEETS 

Information on disease, insect, and mite 

pests that affect Christmas trees in Pennsylvania 

is presented in the following 40 

pest fact sheets. The list of pests is not 

all inclusive. Pests that cause occasional 

or minimal damage or pests that may be 

introduced after printing of the manual 

are not included at this time but may be 

added at a later date. If you are using this 

manual outside of Pennsylvania, you may 

encounter pests that are not included because 

they do not occur in our state. The 

pests have been divided into three categories 

based on the type of injury they cause: 

Needle Discoloration and Injury (pages 

22–71), Shoot and Branch Injury (pages 

72–104), and Stem and Root Injury/Tree 

Mortality (pages 106–121).

NEEDLE DISCOLORATION 

AND INJURY 

The pest fact sheets on pages 23–71 are 

for disease, insect, and mite pests that 

damage needles and result in defoliation, 

discoloration, or distortion. The damage 

can vary from slight yellowing of needles 

to total loss of needles. Tree death is uncommon 

and only occurs if a pest population 

is unmanaged for several seasons. 

NEEDLE DISCOLORATION AND INJURY ................................................................................................................................................................................................................................................ 22

Hosts 

• All species of Christmas trees and 

ornamental conifers 

• Occasionally found on deciduous trees 

and shrubs 

• Host list includes more than 120 species 

of trees and shrubs 

Damage Potential 

• Moderate–high 

Symptoms and Signs 

• Brown spots on foliage 

• Missing needles (current year’s growth) 

• Brown, conelike bags hanging from 

branches 

• Dead branches 

Causes of Similar Symptoms 

• Gypsy moths occasionally defoliate some 

Christmas trees 

• Grass bagworm and snail-cased bagworm 

occasionally found on conifers but do not 

cause damage 

Identifi cation 

Bagworm is a caterpillar that molts into a 

moth in the adult stage. They are easiest to 

identify by the bags they construct as they 

feed. Bags on spruce will look completely 

different from those on arborvitae or honey 

locust because the host plant material is 

incorporated into the bag. Each bag can be 

up to 1½–2½ inches (38–63 millimeters, 

mm) long when the larva is mature. If you 

observe a bag closely, you will see that the 

caterpillar’s shiny black head and fi rst pair of 

legs are exposed as it feeds. The robust body 

of the caterpillar is pale yellow, mottled 

with black, and may be up to 1½ inches 

(38 mm) long when mature. 

Adult bagworms are seldom seen. 

Females have no wings and never emerge 

from the bag they construct in the caterpillar 

stage. Since they do not have wings 

or legs, they resemble maggots. The male 

Calendar of Activities 

Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

moths have two pairs of wings and three 

pairs of legs. Their wings are transparent 

with black borders and their bodies are dark 

and fuzzy. Male bagworms are about ¾ inch 

(15–18 mm) long from head to the tip of 

the abdomen. Their wingspan is approximately 

1 inch (25 mm). 

Biology and Life Cycle 

Bagworms overwinter as eggs inside the bag 

constructed by the female (Fig. 1). In late 

May through mid-June, eggs hatch and the 

larvae crawl out the bottom of this bag. 

They spin down on a thin strand of silk (a 

habit known as “ballooning”). Larvae will 

settle to feed on lower branches or may be 

blown to nearby plants during the ballooning 

stage. When they reach a suitable host, 

the larvae begin to feed and produce silk 

to construct individual bags around their 

bodies (Fig. 2). Plant debris is woven into 

Figure 1. Bagworm eggs inside the female’s 

bag. Courtesy of Sandy Gardosik, PDA 

Jan. Feb. March April May June July Aug. Sept. Oct. Nov. Dec. 

Monitor in late August–September for male moth emergence (see Control Options section). 

Figure 2. Immature 

larvae feeding 

on Douglas-fi r; 

needles show 

symptoms of early 

damage. Courtesy 

of Sandy Gardosik, 

PDA 

BAGWORM 

Thyridopteryx 

ephemeraeformis 

(Haworth) 

Bagworm feeding damage. 

Courtesy of Sandy Gardosik, 

PDA 

BAGWORM ............................................................................................................................................................................................................................................................................................ ..................................................................... 24 23

Figure 3. Immature bagworm 

larva. Courtesy of Sandy 

Gardosik, PDA 

Figure 4. Bagworm larva inside 

open bag. Courtesy of Pennsylvania 

DCNR Forestry Archive, 

Bugwood.org (#5020082) 

Figure 5. Heavy bagworm 

infestation on Colorado blue 

spruce. Courtesy of Rayanne D. 

Lehman, PDA 

Figure 6. Bagworm larval bag 

attached with silk to eastern 

white pine. Courtesy of Cathy 

Thomas, PDA 

each bag to camoufl age and protect the 

larva. The larva feeds through an opening at 

the top of the bag, while the pointed end of 

the bag dangles from the host. Very young 

larvae may carry their bags in a snail-like 

manner for a short time (Fig. 3). 

Only the newly hatched larva and the 

male moth can be found outside a protective 

bag. As the larva grows, it enlarges the 

bag by adding more silk and plant material 

(Fig. 4). The bag is attached to branches 

of the host with loose strands of silk and 

can be moved to get to fresh plant material 

(Fig. 5). After feeding for 8–10 weeks, the 

larva fi rmly attaches the bag to a branch 

with stronger, more durable silk and pupates 

inside the bag (Fig. 6). About 4 weeks later, 

the adult male bagworm emerges. 

After a male moth (Fig. 7) emerges 

through the bottom of the bag, the pupal 

case can occasionally be seen partially 

protruding from the empty bag (Fig. 8). The 

female never leaves her bag but positions 

herself over a small opening in the bottom 

of the bag and begins to emit sex pheromones 

to attract male moths. Mating occurs 

with the male outside the bag and female 

inside. Adults do not feed and die shortly 

after mating. Each female can produce 

300–1,000 eggs before dying. The eggs will 

remain inside the female’s mummifi ed body, 

inside her bag, through the winter. If you 

squeeze a bag containing eggs during the 

winter months, the small, whitish eggs will 

ooze out and may resemble minute tapioca. 

Only one generation of bagworm occurs per 

year in Pennsylvania. 

Monitoring and Management 

Strategies 

Plantation Establishment 

• No recommendations are available 

at this time. 

Preseason 

• Before new growth starts, scout trees for 

brown bags; hand-pick and destroy bags. 

Remove tough silk around the branch to 

prevent girdling as the branch grows. 

• Before mid-May, tag at least one tree 

infested with bagworm to monitor for 

emergence of larvae. 

Figure 7. Adult male bagworm adhered to a 

pheromone-baited sticky trap. Courtesy of 

Tracey Olson, PDA 

Figure 8. Pupal 

case exposed after 

male bagworm 

emergence. 

Courtesy of Sandy 

Gardosik, PDA 

Growing Season 

• Threshold level: At this time, no threshold 

level has been established, but infestations 

on spruce can easily cause economic 

damage. 

• Scout for emerging young larvae on 

lower branches in late May to early June. 

Typically, emerging larvae drop down from 

bags to begin feeding and gradually move 

to the top of the plant as they mature. 

• Growing degree days: Based on observations 

in Pennsylvania, larvae emerge at 

650–750 GDDs. 

• At the end of the season, evaluate results 

and update records. 


Control Options 

Biological 

• Bagworm has several naturally occurring 

insect predators (wasps and hornets) 

and parasitoids (wasps and fl ies) (Fig. 9), 

fungal parasites, and bird predators. No 

species are available for augmentation at 

this time. 

Mechanical 

• Hand-picking and destroying bags anytime 

during growing season or in spring 

before eggs hatch can be very effective in 

eliminating a localized infestation. 

• Remove and burn or fi nely chip severely 

infested trees within and around the 

plantation. 

Biorational 

• Bacillus thuringiensis (Bt) is a bacterium 

that kills specifi c insects, such as caterpillars, 

but is safer for natural enemies. It is 

most effective when used against young 

larvae. Larvae must consume Bt on the 

plant material; therefore, coverage and 

timing are critical. 

• Pheromone traps can be deployed in 

August to trap male moths and decrease 

the number of mated females (Fig. 10). 

These traps contain a chemical that attracts 

males by mimicking the female sex 

pheromone. When males are trapped, 

fewer matings occur and the number of 

overwintering eggs will decrease. 

Chemical 

• Insecticides are most effective when 

applied to young larvae. After fi rst treatment, 

monitor populations to determine 

if second treatment is warranted. 

Next Crop/Prevention 

• Inspect new seedlings for signs of bags; 

remove bags. 

Figure 9. Parasitoid wasp (circled) inserting her 

eggs into the bagworm larval case. Courtesy of 

Sarah Pickel, PDA 

Figure 10. Bagworm pheromone-baited sticky 

trap. Courtesy of Sarah Pickel, PDA 

BAGWORM ............................................................................................................................................................................................................................................................................................. 25

BALSAM TWIG 

APHID 

Mindarus abietinus 

Koch 

Twisted needle symptom. 


Hosts 

• All species of true fi r, especially Fraser and 

balsam 

• Rarely found on some spruce and 

juniper 




• Curled, twisted needles on current year’s 

growth 

• Stunted needles 

• Black sooty mold and presence of stinging 

insects 


• Pesticide use 

• Soft scales 

• Other aphids 


Balsam twig aphids are tiny, soft-bodied 

insects with piercing-sucking mouthparts. 

Most stages are pale bluish green and some 

may have powdery, wax strands adhering to 

the body. The second generation is the only 

one that produces winged forms; all other 

generations have only wingless adults. The 

largest stage, the “stem mother,” occurs in 

the fi rst generation and at maturity is 1 ⁄25– 1 ⁄13 

inch (1–2 mm) long. 

Eggs are small ovals coated with waxy 

rods that have sloughed off from the 

underside of the female. Initially, eggs are 

pale tan, but they darken with age and by 

spring appear to be silvery black. With the 

white, waxy rods and almost black eggs, it is 

relatively easy to spot eggs using a hand lens 

of at least 15X magnifi cation. 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 


The balsam twig aphid has a complex life 

cycle with three to four generations occurring 

per year. Most of the year is spent in 

the silvery egg stage on twigs of the host tree 

(Fig. 1). In early April, prior to bud break, 

the overwintering egg hatches into a small 

nymph (Fig. 2). This nymph feeds for a 

period of time on the underside of last year’s 

needles before molting into the wingless 

stem mother. As buds are swelling and just 

beginning to open, the stem mother moves 

Figure 1. Silvery balsam twig aphid egg at 

the needle’s base on the underside of a twig. 

Courtesy of Rayanne D. Lehman, PDA 

Figure 2. Newly hatched nymph feeding on 

needles and secreting “honeydew.” Courtesy of 

Rayanne D. Lehman, PDA 


NEEDLE DISCOLORATION AND INJURY .......... ................................................................................................................................................................................................................................................ 26

to the bud (Fig. 3) and gives live birth to 

up to 70 second-generation nymphs. No 

mating or eggs are involved; the nymphs are 

clones of the stem mother and capable of 

feeding immediately. 

Figure 3. Stem mother on needle bud. Courtesy 


This second generation is the most 

damaging (Fig. 4). Perfectly synchronized 

with bud break, the nymphs begin to feed 

on the elongating needles (Fig. 5), causing 

distortion and stunting (Fig. 6). Within a 

short time, they have matured and eventually 

produce live, either wingless or winged 

females. If wingless forms are produced, they 

continue to feed on the same tree. Although 

the winged females are weak fl iers, they are 

capable of short fl ights or becoming airborne 

to be carried by air currents to new hosts. 

When the winged females reach the 

new host or new stem on same tree, they 

produce both wingless females and males 

(Fig. 7). This last generation is the only 

one with males that mate with the females. 

Mated females deposit one or two overwintering 

eggs close to the buds before dying. 

This is also the only time in the life cycle 

that eggs are produced. 


Strategies 


• Avoid planting at lower elevations and on 

slopes where aphid populations tend to be 

higher. 

• Plant species that have delayed bud break. 

Balsam fi r tends to break bud early and is 

most severely infested; Fraser is next to 

break bud; Canaan generally breaks bud 

last. 

• Do not buy infested trees or nursery stock. 

Preseason 

• Between July and March, assess damage 

levels from the previous year’s infestation. 

It is most important to control this pest 

during the last 2 years before harvest in 

order to produce 2 years of good, straight 

growth. On young trees, excessive damage 

will stunt overall growth. 

• Do not apply a nitrogen fertilizer before 

bud break to a previously infested area. 

This tends to increase aphid populations. 

• Look for overwintering eggs and tag trees 

to monitor for egg hatch by the end of 

March. 

• Growing degree days: Egg hatch occurs at 

30–100 GDDs. 

• Begin monitoring for egg hatch by April 

1 in southern counties. Using a hand 

lens, examine the underside of last year’s 

needles for single, blue-green nymphs. 

These nymphs are usually within 2 inches 

of the bud and often have a large droplet 

of clear honeydew on their tail end. 

• Encourage natural predators and parasitoids. 


• Scout for aphids by beating 10 inches of 

foliage over a piece of dark paper. Select 

15 trees per acre (similar age, size, and 

location) and sample two sides of each 

tree. Count aphids and predators found. In 

April or early May, it may be necessary to 

apply chemical control if more than three 

aphids per tree are found. 

• As bud break begins, look at the opening 

buds to fi nd the stem mothers and secondgeneration 

nymphs. 

• When vegetative cones are produced on 

trees, break open the cones and look for 

nymphs that have migrated here to feed. 

• Observe presence of lady beetles since 

several species are important predators. 

• Observe presence of stinging insects as 

new growth elongates. The bees and wasps 

are attracted to the honeydew produced 

by the feeding aphids. 

• At the end of the season, update records 

and evaluate results. 

Figure 4. Balsam 

twig aphid nymphs. 

Courtesy of 

Rayanne D. Lehman, 

PDA 

Figure 5. Nymphs (circled) feeding 

in a barely opened bud. 


PDA 

Figure 6. Needle twisting or 

kinking evident of balsam twig 

aphid damage. Courtesy of 


Figure 7. Winged balsam twig 

aphid female. Courtesy of 


BALSAM TWIG APHID ............................................................................................................................................................................................................................................................................ 27

Figure 8. Larva of multicolored 

Asian lady beetle. Courtesy of 



Biological 

• Balsam twig aphids have numerous 

naturally occurring predators, including 

yellow jackets, lacewings, earwigs, lady 

beetles and their larvae (Fig. 8), assassin 

bugs, ants, big-eyed bugs, predatory thrips, 

hover fl y larvae, syrphid fl ies, and predaceous 

midges. The introduced multicolored 

Asian lady beetle is an excellent 

aphid predator in the adult and larval 

stage. Parasitoids of balsam twig aphids are 

Aphidius wasps. Refer to Appendix B: Biological 

Controls Photo Chart for pictures. 

Mechanical 

• Pick and destroy young cones in spring 

during aphid activity. Since pesticides will 

not reach the aphids inside cones, this 

practice can be time consuming but very 

benefi cial and will positively affect the 

aesthetics of the tree. 

Biorational 


at this time. 

Chemical 

• All pesticide intervention should occur 

after egg hatch but before bud break; 

after bud break, aphids are protected by 

new growth and some damage has already 

occurred. Dormant oil will kill aphids but 

will have limited effect on overwintering 

eggs and often causes damage to elongating 

needles. 


• Purchase and plant pest-free nursery stock 

from a reputable company. 


Hosts 

• Douglas-fi r 

• Alternate hosts: Colorado blue spruce and 

occasionally other spruces (see Shoot and 

Branch Injury section for Cooley spruce 

gall adelgid on spruce) 


• Moderate 


• Yellow spots on the needles 

• Needles with bends or crooks 

• Small, white, cottony balls on the 

underside of needles 

• Premature needle drop 


• Rhabdocline needle cast 

• Douglas-fi r needle midge 


The Cooley adelgid is a small, soft-bodied 

insect with sucking mouthparts. Both 

winged and wingless forms exist, but the 

winged insect is not often seen. The easiest 

method of identifi cation is the characteristic 

damage. On Douglas-fi r, Cooley adelgid 

does not cause galls, as it does on spruce. 

Instead, it causes pale or yellow spots on the 

top of the needle. Frequently, the needle 

will have a bend or crook at this site. These 

spots or bends indicate feeding sites for the 

nymphs, which will be found on the underside 

of the needle. The crawlers and early 

nymphs are dark brown to black, fl at, and 

have ridges across their bodies. They can 

be seen using a hand lens with at least 15X 

magnifi cation. White, cottony masses may 

also be found on the underside of needles at 

certain times of year. 

In early spring, examine the underside of 

needles on Douglas-fi r. The egg masses will 

be beneath a waxy mass on needles of outer 

growth. At bud break, look at the base of 

elongating shoots for minute, dark nymphs 

feeding on the new needles. 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 


The life cycle of Cooley adelgid is 

complex and involves distinctly different 

forms on Colorado spruce and Douglasfi 

r. The pest can alternate between the 

two hosts or produce continual generations 

on either. Both winged and 

wingless forms are found; all adults on 

Douglas-fi r are female. 

On Douglas-fi r, exposed nymphs 

overwinter on the underside of the 

needles (Fig. 1). In early spring, these 

nymphs begin to feed and produce 

their waxy protective threads. Initially, 

the white wax is just a fringe around 

the dark body (Fig. 2), but by the time 

they reach maturity, the entire body is 

engulfed (Figs. 3 and 4). 

Figure 1. 

Overwintering 

nymphs. Courtesy 


PDA 

Figure 2. Nymph 

with waxy fringe 

beginning to form 

around the body. 


Gardosik, PDA 

Figure 3. Nymph 

covered with white, 

waxy fringe. 


Gardosik, PDA 


Spray must occur before nymphs wax over. 

Figure 4. Nymphs 

covered with white, 

waxy fringe. 

Courtesy of Brian 

Schildt, PDA 

COOLEY 

SPRUCE GALL 

ADELGID ON 

DOUGLAS-FIR 

Adelges cooleyi 

(Gillette) 

White, waxy tufting of 

Cooley spruce gall adelgid. 


PDA 

COOLEY SPRUCE GALL ADELGID ON DOUGLAS-FIR ........................................................................................................................................................................................................................... ..................................................................... 29

Figure 5. Cooley eggs laid 

under the waxy fringe of the 

mature female. Courtesy of 

Sandy Gardosik, PDA 

Figure 6. Cooley nymphs 

feeding on new growth of 

Douglas-fi r, resulting in needle 

kinking. Courtesy of PDA 

Figure 7. Kinked needles and 

chlorosis caused by nymph 

feeding. Courtesy of 


Two types of females result from these 

overwintering forms: wingless females that 

will remain on Douglas-fi r and winged 

females that can fl y to spruce and complete 

a nondamaging generation. The wingless 

females produce a cluster of up to 100 eggs 

(Fig. 5), which begin to hatch as trees break 

bud. Nymphs crawl into the opening buds 

and feed on the elongating needles (Fig. 6). 

Chlorotic spots and bending of the needles 

result from the nymphs feeding, but no gall 

results. (Fig. 7). Subsequent generations 

occur on Douglas-fi r through the season, but 

the fi rst generation is the most damaging. 


Strategies 


• When planting, separate Colorado blue 

spruce and Douglas-fi r trees. This will not 

eliminate problems, but it may help lessen 

the severity. 

• Plant resistant/tolerant Douglas-fi r 

varieties. 

• Remove any mature Colorado blue spruce 

or Douglas-fi r that may be sources of 

infestation. 

Preseason 

• Scout for overwintering nymphs on the 

underside of needles. 

• Examine the undersides of needles on 

inner branches as well as last year’s 

growth. 


• Growing degree days: The recommended 

control period against nymphs is 22–81 

GDDs in the spring (before nymphs wax 

over) and 2,800–3,000 GDDs in the fall. 

• Threshold level: No threshold has been 

established. Control for 2 years before 

harvest to have damage-free needles. 




Biological 

• Encourage natural predators such as 

lacewings, assassin bugs, and lady beetles. 

Mechanical 


at this time. 

Biorational 


at this time. 

Chemical 

• Dormant oil: Overwintering nymphs can 

be controlled by applying dormant oil 

before new growth starts in early spring or 

in late fall after fi rst frost when trees are 

not actively growing. 

• Spring insecticide: The fi rst application 

should occur after nymphs/immature 

females begin to swell but before they 

produce white, waxy threads to cover 

themselves. A second application in 7–10 

days may be needed but must occur before 

bud break. 

• Fall insecticide: A single spray should be 

applied in late September or October to 

control the exposed nymphs and immature 

females before overwintering. 

• Insecticide applications are not recommended 

during the growing season since 

most stages will be protected under waxy 

threads or inside galls. 





Hosts 

• True fi rs, especially Fraser, Canaan, and 

balsam 

• All other conifer species 




• Damage to denser, lower, and inner 

branches of the tree 

• Mottled yellowing on needle tops 



• Elongate hemlock scale 

• Spruce spider mite 


Cryptomeria scale is an armored scale found 

on the underside of needles. Observed with 

a hand lens, the scale resembles a fried 

egg. The yellow, soft-bodied scale insect is 

protected beneath a whitish-gray, oval cover. 

Yellow, cast skins of the immature scales can 

be seen centrally under the translucent scale 

cover. The scale covering is the structure 

most frequently observed and will easily 

slough off when rubbed with your fi nger. 

When the scale covering is removed, it 

frequently leaves a solid green ring on the 

white stomata on the needle. The female 

covering is about 1 ⁄20 inch (1.0–1.5 mm) 

long and the immature male covering is 

slightly smaller. Underneath the covering, 

the adult female scale is a yellow, fl attened 

oval that lacks legs or a distinct head. Prying 

the insect from the needle may reveal the 

extremely long, thin mouthparts inserted 

in the needle. Adult male scales are winged 

and rarely seen. Eggs deposited under the 

female covering are oval and yellow. The 

pale yellow, lozenge-shaped crawlers are 

about 1 ⁄100 inch long (0.25 mm) and visible 

to the naked eye after they have moved from 

under the female cover. 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 


All stages of Cryptomeria scale are found on 

the underside of needles. The crawlers and 

males are the only stages capable of moving 

about on the needles. All other immature 

stages and the females are sedentary, remaining 

on the site where they fi rst inserted their 

piercing-sucking mouthparts to feed (Fig. 1). 

This feeding causes chlorotic spotting on the 

needles (Fig. 2), which may give an infested 

tree a yellow appearance beginning at its 

base (Fig. 3). 

Figure 1. 

Heavy 

infestation 

of mature 

Cryptomeria 

scales. 

Courtesy of 

Sandy 

Gardosik, PDA 

Figure 2. Chlorotic damage from scale feeding. 


Figure 3. 

Yellowed 

trees with 

Cryptomeria 

scale damage. 

Courtesy of 

Cathy Thomas, 

PDA 


CRYPTOMERIA 

SCALE 

Aspidiotus cryptomeriae 

Kuwana 

Chlorotic damage from 

Cryptomeria scale feeding on 

the undersides of needles. 


PDA 

CRYPTOMERIA SCALE ........................................................................................................................................................................................................................................................................... ..................................................................... 29 31

Figure 4. Adult male Cryptomeria 

scale emerged from its scale 

covering. Courtesy of Sandy 

Gardosik, PDA 

Figure 5. Adult female scale 

without its scale covering and 

other mature scales. Courtesy 

of Sandy Gardosik, PDA 

Figure 6. Exposed female scale 

with eggs. Courtesy of Sandy 

Gardosik, PDA 

After overwintering as immatures, the 

scales resume feeding and mature by late 

April. Males have an additional nonfeeding 

pupal stage before the winged male 

(Fig. 4) emerges and fl ies to a female (Fig. 

5). With the female under her protective 

cover, mating occurs and the male dies. 

Females deposit approximately 40 yellowish 

eggs beneath the cover before dying by late 

May or early June (Fig. 6). Crawlers emerge 

about 2 weeks after the eggs are deposited. 

The crawler emergence period may extend 

for 6–7 weeks but generally peaks about 

2–3 weeks after eggs are laid. 

Crawlers are relatively fragile and desiccate 

quickly (Fig. 7). They can crawl short 

distances but must settle and begin to feed 

within a day of emerging from the egg. As 

they settle and begin to feed, they create 

the waxy covering to protect themselves 

from environmental conditions (Fig. 8). It 

is not uncommon to fi nd a settled crawler 

beginning to develop its protective covering 

under the dead female. Several layers 

from different generations may develop with 

continual infestation. 

Figure 7. 

Crawler 

(circled), 

settled crawlers, 

female, 

and eggs. 

Courtesy of 

Sandy 

Gardosik, PDA 

Figure 8. 

Settled scale 

crawlers. 

Courtesy of 

Sandy 

Gardosik, PDA 

In summer, the stages repeat and by late 

July or early August, eggs of the second generation 

are present. The second-generation 

crawlers begin to emerge in mid-August, 

but some crawlers may be present into 

October. Two generations occur, but not all 

individuals mature at the same time, so the 

generations are not as distinct as some other 

multiple-generation pests. 


Strategies 


• Plant tree varieties that are less susceptible 

to Cryptomeria scale infestations. 

• Weed management in rows is important 

since it permits easier scouting and more 

thorough coverage if chemical controls 

are needed. 

Preseason 

• Scout for scale insects on the underside of 

needles, particularly those on the inside of 

bottom branches. Look for trees that exhibit 

yellow mottling on the upper surface 

of the needles (Fig. 9). If scale insects are 

found, verify that it is Cryptomeria scale 

by comparing the description with that 

of elongate hemlock scale. Male elongate 

hemlock scale coverings are white. 

• Scouting on an overcast day will allow 

symptoms to stand out. 

• Place sticky cards on branches showing 

symptoms to trap adult male scale insects 

(Fig. 10). Their emergence signals that 

eggs will soon follow. 

Figure 9. Lifting lower tree branches to scout 

for Cryptomeria scale. Courtesy of Cathy 

Thomas, PDA 

Figure 10. Sticky cards for monitoring adult 

male scale emergence. Courtesy of Cathy 

Thomas, PDA 


• If only a few trees are infested, remove 

and destroy the trees before bud break. 

• If the block is generally infested, tag 

several trees that can be easily observed 

for crawler emergence. 


• Growing degree days: 

— First-generation crawlers begin to 

emerge at 600–800 GDDs. 

— Second-generation crawlers begin to 

emerge at 1,750–2,130 GDDs. 

• Threshold level: Currently, no threshold 

level has been established. Cryptomeria 

scale can cause damage with just a few 

scales per needle. 

• Scout tagged trees for egg deposition in 

late May, and then scout daily for fi rst 

crawlers. Repeat scouting regime in late 

summer for second generation. 




Biological 

• Encourage naturally occurring parasitoids 

and predators. 

• Do not use a broad-spectrum insecticide 

that will kill benefi cial insects. 

Mechanical 

• Remove and destroy heavily infested trees 

before bud break. Wrap trees in a tarp/ 

plastic when dragging them through the 

fi eld to prevent transferring scales to other 

trees. 

• Clean mower blades or tractors when 

moving them from an infected fi eld to an 

uninfected fi eld. 

• Butt-prune infested trees to remove the 

most heavily infested lower branches. 

Biorational 


at this time. 

Chemical 

• Apply dormant oil in spring before bud 

break. Thorough coverage of underside of 

lower branches is important. 

• For fi rst- or second-generation control, 

apply an appropriate spray targeting 

the crawlers after they are fi rst are seen. 

Repeat sprays every 7–10 days for control; 

up to four sprays may be required. Evaluate 

infested branches for presence of new 

crawlers after each spray to determine if 

another spray is necessary. Check needles 

from two to three seasons back for crawlers. 


• Purchase and plant scale-free nursery 

stock from a reputable company. 

• Remove heavily infested trees from a fi eld 

of new plantings as well as infected lower 

branches left behind after tree cutting. 

CRYPTOMERIA SCALE ............................................................................................................................................................................................................................................................................ 33

CYCLANEUSMA 

NEEDLE CAST 

Cyclaneusma minus 

(Butin) DiCosmo, 

Peredo, and Minter 

(syn. Naemacyclus 

minor Butin) 

Yellow-brown needles on 

infected tree. Courtesy of 

USDA Forest Service, 

Forest Health Protection, St. 

Paul Archive, Bugwood.org 

(#5050051) 

Hosts 

• Pines, esp. Scotch 

• Also reported on Austrian, Ponderosa, 

Mugo, and Monterey pines 


• Moderate 


September Through November 

• Light green spots that eventually become 

yellow on needles 2 years old and older 

• Yellow needles with transverse brown 

bands 

October Through May 

• Symptomatic needles anywhere on the 

tree or cast to the ground 

• Off-white fruiting bodies that develop 

on yellow needles and swell with wet 

weather, making them easier to see 


• Aphid feeding 

• Other needle casts such as Lophodermium, 

Lophodermella, and Dothistroma 

• Environmental stresses 

• Air pollution 

• Fall needle drop 

• Winter injury 

• Pine needle scale 


Cyclaneusma needle cast is commonly referred 

to as the “fall yellower” of Scotch pines 

because of the timing and appearance of 

initial symptoms. Symptoms appear on older, 

interior needles from 10 to 15 months after 

infection occurs. In late summer or early fall, 

infected needles will have light green to yellow 

spots reminiscent of aphid or scale insect 

feeding. Gradually, these yellow spots enlarge 

to form yellow bands, eventually causing the 

entire needle to turn yellow. By winter, the 

needles are light tan, and brown transverse 

bands become visible. Infected needles remain 

on the tree through the winter. Fruiting 

bodies form about one month after symptoms 

appear. Because they are off-white in color 

and similar to the surrounding diseased tissue, 

they are diffi cult to detect. These structures 

become more obvious during periods of moist 

weather when they swell and split the epidermis 

of the needle, causing it to look like two 

open doors. 

Because many needle casts are similar, 

laboratory diagnosis may be warranted for 

proper identifi cation before applying a 

fungicide. 


Cyclaneusma needle cast causes many pine 

species to prematurely defoliate. Some 

research suggests that this fungus may be an 

opportunistic pathogen that causes disease 

on plants that are under environmental 

stress. Four periods of infection occur for 

this fungal disease. The fi rst is from mid-July 

through August, when the current year’s 

needles may become infected. September 

through November and late November 

through early December are the second and 

third infection periods. In Pennsylvania, 

these three periods account for about half of 

the new infections. The remaining 50 percent 

of infections occur from April through 

June, when both mature needles and new 

growth are susceptible. 

Spores are dispersed by the wind as they 

are released by fruiting bodies. If proper 

moisture is present, the fungus will enter 

the needle through stomata. About the 

time needles become discolored with brown 

bands, fruiting bodies will develop beneath 

the surface of the needle (Fig. 1). They begin 

to swell and the epidermis of the needle 

opens with a longitudinal fi ssure (Fig. 2). 

When moist or wet, the fruiting bodies 

enlarge and forcibly discharge spores. 

Figure 1. Yellow-brown bands on infected 

Scotch pine needles. Courtesy of Tracey Olson, 

PDA 

Figure 2. Swollen, light brown fruiting bodies 

that will rupture the surface of the needle. 

Courtesy of Joseph O’Brien, USDA Forest 

Service, Bugwood.org (#5050047) 


Disease Cycle Calendar (Single Year’s Growth of Needles) 

May J J A S O N D Jan. F M A M J J A S O N D Jan. F M A M J J A S O N D 

Infection 

Symptoms 

Casting 

First Year Second Year Third Year 

Bud Break 

The heavier the shading, the more intense the infection/symptom/casting. 

Throughout the year, spores are released 

from infected needles that are attached or 

have fallen from the tree (Fig. 3) when the 

temperature is above freezing and there is 

moisture on the needle. Infected needles may 

remain on the tree or be cast anytime after 

they begin exhibiting symptoms (Fig. 4). 

Figure 3. Cast needles may continue to 

sporulate. Courtesy of Joseph O’Brien, USDA 

Forest Service, Bugwood.org (#5050046) 

Figure 4. Infected needles still attached to the 

tree. Courtesy of Joseph O’Brien, USDA Forest 



Strategies 


• Choose a site that will promote drying 

of trees (southern slope, good drainage, 

nonshaded). 

• Provide adequate spacing between trees to 

allow for air circulation. 

• Plant stock that shows resistance or tolerance 

to the disease; use seed stock that 

exhibits genetic resistance. 

Preseason 

• Maintain proper nutrient and water levels 

to keep trees healthy and vigorous. 

• Scout trees in susceptible areas for any 

symptoms of the disease (a hand lens may 

be required). 

• Maintain proper weed control; do not 

allow weeds to grow up under or between 

the trees. 


• Threshold level: If more than 20 percent 

of sampled trees have signs of the disease, 

consider treating the entire plantation. 



CYCLANEUSMA NEEDLE CAST .............................................................................................................................................................................................................................................................. 35


Biological 


at this time. 

Mechanical 

• Remove and destroy severely infected 

trees. 

• Remove dead needles beneath infected 

trees. 

• Examine needles around stumps for symptoms; 

remove infected needles if present. 

• Shear nondiseased trees prior to diseased 

trees to minimize spread of the fungus. 

Biorational 


at this time. 

Chemical 

• Fungicide application: In Pennsylvania, 

fi ve spray applications of an appropriate 

fungicide effectively treat the disease; 

spray in late March, early May, mid-June, 

mid-August, and early October. 


• Buy disease-free and/or resistant stock 

from a reputable nursery. 

• Scout in late fall for new trees just starting 

to show signs of infection. 

• Avoid planting next to old Scotch pine 

stands that may harbor disease. 


Hosts 





Summer Through Fall 

• Swollen areas on current year’s needles; 

needle may bend at gall 

• Pale or yellow spot(s) on both sides of 

needle; needle may turn purple or brown 

in fall 

• Premature needle loss 



• Cooley adelgid 


The Douglas-fi r needle midge is a tiny, 

orange-colored fl y, approximately 1 ⁄8 inch 

(5 mm) long. During the emergence period 

in spring, look for minute, orange fl ies resting 

on the tips of needles. Females can be 

distinguished by a very long ovipositor with 

which they probe between bud scales and 

into partially opened buds. This elongate 

ovipositor enables the female to deposit 

the long, narrow, orange-colored eggs in 

protected areas. A hand lens is needed to 

view the eggs. 

A white, legless maggot lacking a distinct 

head hatches from each egg and bores 

directly into the needle. In response to the 

maggot’s feeding, the needle produces excessive 

tissue, resulting in one or more swollen 

areas on the needle. There may be a minute, 

orange spot visible on the underside of the 

needle at the larval entrance site. When 

mature galls are carefully sliced open, a pale 

maggot may be found, generally under the 

entrance spot. At maturity, the maggot is 

about 1 ⁄8 inch (5 mm) long. After the maggot 

has dropped out of the needle to pupate 


in the ground, an irregular, three-sided exit 

hole can be seen in the epidermis on the 

underside of the affected needle. In fall and 

winter, damaged needles drop from the tree 

or break at the gall location, making winter 

scouting more diffi cult. 


Douglas-fi r needle midge overwinters as a 

mature larva in soil beneath an infested 

tree. In March–April, it pupates in the soil 

and the adult midge emerges (Fig. 1). The 

emergence period is generally from mid- 

April through early May and depends on the 

weather conditions at each site. Rain and 

cool temperatures delay emergence. Mating 

and oviposition occur immediately after 

emergence and adults die shortly thereafter 

(Fig. 2). The peak emergence period lasts 

about 7–10 days. 

Figure 1. Female 

Douglas-fi r needle 

midge. Courtesy 


PDA 

Figure 2. Adult midge (circled) resting on a 

Douglas-fi r bud. Courtesy of Sandy Gardosik, 

PDA 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

*Place emergence traps during the monitoring time. Control measures occur based on trap catch. 

DOUGLAS-FIR 

NEEDLE MIDGE 

Contarinia pseudotsugae 

Condrashoff 

Kinking caused by 

Douglas-fi r needle midge 

larvae. Courtesy of Tracey 

Olson, PDA 

DOUGLAS-FIR NEEDLE MIDGE .............................................................................................................................................................................................................................................................. ..................................................................... 29 37

Figure 3. Eggs found tucked 

in the needle bud before buds 

are fully broken. Courtesy of 


Figure 4. Young larva found 

inside a needle. Courtesy of 


Figure 5. Early evidence of galls; 

needles are beginning to kink 

(late July). Courtesy of Tracey 

Olson, PDA 

Figure 6. Advanced gall 

symptoms (November). 


Figure 7. Mature larva still in 

the needle (December). 


Eggs (Fig. 3) hatch within a few days and 

larvae (Fig. 4) bore directly into the young 

needles to feed throughout the summer. As 

the larvae feed and grow, galls form in the 

needles and cause them to kink (Fig. 5). 

By late summer, the galls will begin to 

turn yellow and then brown (Fig. 6). In late 

fall, the full-grown larvae (Fig. 7) exit the 

needles and drop to the ground to overwinter, 

leaving a distinct triangular exit hole 

in the underside of the needle (Fig. 8). 

A single generation occurs each year. 


Strategies 


• Plant late breaking tree varieties to 

decrease midge damage. 

• Remove overgrown Douglas-fi rs from the 

perimeter of the block. Damage is often 

more severe on older trees. 

Preseason 

• Place emergence traps under the north 

side of previously infested trees by April 

1 or before the daily temperature reaches 

60°F for several days (Fig. 9). See Appendix 

E: Insect Trap Use and Construction 

for information on trap construction. 

• Place at least three traps per fi eld and 

check every day until the fi rst midge appears. 

Record the number of midges in the 

trap jar before emptying and replacing the 

jar. Continue monitoring until no midges 

are present for several days. 


• Growing degree days: The adult midges 

emerge from the soil beneath trees at 

200–400 GDDs (based on midge population 

monitoring in Pennsylvania). 

• Threshold level: No established threshold 

level exists for this pest. 

• Scout in August for infestation. Mark 

trees that exhibit symptoms to aid with 

trap placement and distribution of control 

chemical next spring. 



Figure 8. Needle 

showing larva 

exit hole. 

Courtesy of 

Tracey Olson, 

PDA 


Biological 

• Encourage and protect natural predators 

such as tiny parasitic chalcid wasps. 

Ill-timed chemical application is not 

effective and may kill natural enemies. 

Mechanical 

• Remove heavily infested trees in early fall 

before larvae exit the needles. 

Biorational 


at this time. 

Chemical 

• Base application on collection of adults 

in emergence trap. Chemicals will not be 

effective against eggs, only adults. First 

application should be made within a day 

of collection of the fi rst midge. 

• Consider a second application in 2 weeks 

if adults are continuing to emerge. 




Figure 9. Emergence trap placed under a 

previously infected tree. Courtesy of Sandy 

Gardosik, PDA 


Hosts 

• Fir (Abies spp.) 

• Hemlock (Tsuga spp.) 

• Occasionally found on spruce (Picea spp.), 

Douglas-fi r (Pseudotsuga menziesii), and 

pines (Pinus spp.) 


• Low–high 


• Yellowing of needles on the interior of the 

lower branches; damage moves upward as 

population increases 

• Scale coverings on the underside of the 

needles 

• Premature needle drop; eventual branch 

and limb dieback and death of tree with 

severe infestations 

• Tree may appear fl ocked 



• Cryptomeria scale 


Elongate hemlock scale—sometimes 

referred to as Fiorinia scale—is an armored 

scale found more prevalently in southeastern 

Pennsylvania. Translucent, oval eggs 

occur beneath the scale covering of the 

female. The pale yellow crawlers are about 

1 ⁄250 inch (0.1 mm) long and have six short 

legs. When they settle to feed, the crawlers 

lose their legs and excrete an amber-colored, 

oval covering. Male and female scales develop 

differently and have different protective 

coverings. Immature males are about 

1 ⁄20 inch (1.0–1.5 mm) long and produce a 

whitened, waxy covering. Longer threads of 

wax are occasionally present and may give 

the covering a fuzzy appearance. Immature 

females are longer at 1 ⁄14 inch (1.5–2.0 mm) 

and produce a yellowish to orange-brown, 

parallel-sided covering. At maturity, the 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

delicate, light brown, male scales emerge 

and fl y to the sessile, wingless females. 

They may be mistaken for parasitoids as 

they crawl over the female coverings prior 

to mating. 


Elongate hemlock scale overwinters in 

several developmental stages in Pennsylvania 

and matures in spring (Figs. 1 and 2). 

In southern and Mid-Atlantic states, two 

overlapping generations have been reported, 

while in states in the Northeast, a single 

generation is known. In Pennsylvania, 

reproduction is staggered, so crawlers are 

present throughout the growing season. 


Apply dormant oil prior to bud break. 

Figure 1. Heavy infestation of elongate hemlock 

scale. Courtesy of PDA 

Figure 2. Female (brown) and male (white) 

scales. Courtesy of Sandy Gardosik, PDA 

ELONGATE 

HEMLOCK 

SCALE 

Fiorinia externa Ferris 

Chlorotic damage from 

elongate hemlock scale 

feeding on the undersides of 

needles. Courtesy of Sandy 

Gardosik, PDA 

ELONGATE HEMLOCK SCALE ................................................................................................................................................................................................................................................................ ..................................................................... 29 39

Figure 3. Female scales and 

mobile yellow crawlers on the 

needles. Courtesy of Sandy 

Gardosik, PDA 

Figure 4. Chlorotic scale feeding 

damage on grand fi r. Courtesy 

of Cathy Thomas, PDA 

Figure 5. Female elongate 

hemlock scale. Courtesy of 


Figure 6. Immature male 

elongate hemlock scales. 


PDA 

The fi rst egg hatch generally starts in 

late May or early June. Mobile crawlers 

move from under the female’s covering to 

the underside of needles (Fig. 3) and begin 

to feed by inserting their piercing-sucking 

mouthparts. This feeding causes chlorotic, 

or yellow, spotting on the upper needle 

surface (Fig. 4). At the same time, the 

crawlers begin to secrete a translucent, 

waxy covering. Mature females are present 

about 6–8 weeks after emerging (Fig. 5). 

Male scales molt an additional time before 

the winged stage emerges (Figs. 6 and 7). 

Following mating, the males die and the 

female begins to produce about 12–16 eggs 

under her armored covering (Fig. 8) As a 

result of the various overwintering stages in 

Pennsylvania, crawlers emerge throughout 

the growing season, making repeated 

control efforts a necessity. 

Figure 7. Adult male elongate hemlock 

scale. Courtesy of Jim Stimmel, PDA 

Figure 8. Eggs inside the overturned female 

elongate hemlock scale casing. Courtesy of 



Strategies 


• Plant tree species that are not susceptible 

to elongate hemlock scale. 

• Properly space trees when planting to 

minimize infestation from tree to tree and 

enable thorough coverage if chemical 

controls are used. 

• Weed management is important to expose 

lower branches for ease of detection and 

effective control. 

• Remove and destroy any mature host trees 

that may serve as source of infestation. 

(Canada hemlock is a common host in 

Pennsylvania.) 

Preseason 

• Limit the use of nitrogen fertilizers; nitrogen 

enhances survival and developmental 

rate of scales. 

• Scout for elongate hemlock scale on the 

underside of needles. Note: Sometimes 

a combined population of Cryptomeria 

scale, balsam woolly adelgid, and hemlock 

scale can be found on the same trees. 

• If only a few infested trees are found, 

removing and destroying them before bud 

break may prevent a serious infestation 

from developing. 

• If numerous infested trees are located, tag 

a few to use for observations of crawler 

emergence. 

• Place sticky cards on branches showing 

symptoms to trap adult male scale insects 

(Fig. 9). Their emergence signals that eggs 

will soon follow. 

Figure 9. Yellow sticky cards used to attract 

the emerging male elongate hemlock scales. 




• Starting in mid-May, scout tagged trees 

every few days to check for crawler 

emergence. 

• Growing degree days: Crawlers are active 

from 360 to 700 GDDs. 

• Threshold level: No specifi c threshold 

level exists at this time. However, dieback 

of major limbs tends to occur by the time 

the population reaches ten scales per 

needle. 

• Continue scouting for crawlers every 

few weeks to determine the need for 

additional controls. 




Biological 

• Encourage natural predators such as 

lady beetles, parasitic wasps, and several 

lacewing species. 

• Avoid applications of broad-spectrum 

insecticides that will kill natural 

predators. 

Mechanical 


before bud break. Wrap trees in a tarp/ 

plastic when dragging them through the 

fi eld to prevent transferring scales to other 

trees. 

• Clean mower blades or tractors when 

moving them from an infected fi eld to an 

uninfected fi eld. 

• Butt-prune infested trees to remove the 

most heavily infested lower branches. 

Biorational 


at this time. 

Chemical 

• Horticultural oil may be applied during 

the growing season. Use lower rate to 

avoid damage to new growth or wait to 

apply until new growth has hardened off. 

• When fi rst crawlers are active, apply a 

systemic insecticide spray three to four 

times over a 12-week period (three sprays 

four weeks apart or four sprays three weeks 

apart). Do not apply more than four times 

per season. 

• Evaluate each application to determine 

the need for subsequent sprays. 




ELONGATE HEMLOCK SCALE ................................................................................................................................................................................................................................................................. 41

ERIOPHYID 

RUST AND 

SHEATH MITES 

Nalepella and Setoptus 

spp. 

Lower branch showing rusty 

feeding damage of the 

spruce rust mite. Courtesy of 


Hosts 

• Spruce 

• Fir 

• Eastern white pine 

• Scotch pine 

• Hemlock 


• Moderate 


Rust Mites (Nalepella spp.) 

• Chlorotic or rusty-brown-appearing 

needles 

• Silvered, dusty, or bleached-out 

appearance on Colorado blue spruce 

• Premature needle drop may occur 

Sheath Mites (Setoptus spp.) 

• Stunted/chlorotic needles; premature 

needle drop 

• Silvering or graying of needles, 

particularly on south side of tree 



• Environmental stress 

• Nutrient imbalances 


Eriophyid mites are distinctly different from 

the typical spider mite. They are extremely 

small and only have two pairs of legs in all 

active stages. The body is wedge or carrot 

shaped and light tan or pinkish (Figs. 1–3). 

Figure 1. Spruce rust mites 

in new growth. Courtesy of 



Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

Figure 2. Hemlock rust mites 

moving along the needle. 


PDA 

Three species of rust mites in the genus 

Nalepella are found on Christmas trees in 

Pennsylvania. Hemlock rust mite, Nalepella 

tsugifolia Keifer, is the predominant species 

on fi r and hemlock. Another species, Nalepella 

octonema Keifer, is found occasionally 

on fi r. The third species, dubbed the spruce 

rust mite (Nalepella halourga Keifer), is 

restricted to spruce. Identifi cation of species 

requires examination under a very highpowered 

microscope. However, because the 

biology, life history, and damage of the three 

species are the same, confi rmation of a rust 

mite infestation is all that is needed. 

Two species of pine sheath mites in the 

genus Setoptus are found in Pennsylvania. 

White pine sheath mite, Setoptus strobacus 

Keifer, can be found in the needle sheath and 

on the needles of eastern white pine. Scotch 

pine sheath mite, Setoptus jonesi Keifer, has 

the same habits but on Scotch pine. 

To scout for eriophyid mites, use a 

15–20X hand lens. Select a lateral branch 

4–5 feet (1.23–1.52 m) from the ground on 

the southern-facing side of the tree. For 

rust mites, turn the branch over to see the 

underside of the needles and look for mites 

and eggs or cast skins. To locate sheath 

mites, look inside the needle sheath at the 

base of the needles. The mites are slow 

moving, and the anterior end is the widest 

portion of the body. You may see two pairs 

of short legs. 

Eggs are round and about the size of the 

needle stomata. During the growing season, 

eggs are generally found in the same location 

as the adults and are off-white or tan. Overwintering 

eggs will be found on the lower 

Figure 3. Wedge-shaped 

eriophyid mites (note four 

legs). Courtesy of Sandy 

Gardosik, PDA 



third of the needle (closer to the needle base 

on fi r) and, like the adults, on the underside 

of the needle (Fig. 4). These eggs range in 

color from clear to salmon to tan and can 

be deposited in clusters of 20 or more (Fig. 

5). To see these eggs, pull needles one by 

one and examine the base. Overwintering 

sheath mite eggs are located on the exposed 

base of needle bundles. During the growing 

season, these eggs will be found in the 

needle sheath and can be located by gently 

pulling needle bundles from the sheath. 

Figure 4. Overwintering 

rust 

mite eggs laid 

near the base 

of the needle. 

Courtesy of 

Sandy 

Gardosik, PDA 

Figure 5. 

Overwintering 

rust mite eggs 

laid in clusters 

of 20 or more. 

Courtesy of 

Rayanne D. 

Lehman, PDA 

Another method of detecting eriophyid 

mites is to look for cast skins on the needles. 

These appear as minute, white strands and 

offer better contrast to the needles than the 

adults. A hand lens is needed to look for 

cast skins (Figs. 6 and 7). If this method is 

employed, an attempt to fi nd adults or eggs 

should follow to confi rm the infestation is 

active. 

Figure 6. 

Threadlike 

cast skins of 

eriophyid 

mites. 

Courtesy of 

Rayanne D. 

Lehman, PDA 

Figure 7. 

Cast skins 

giving the 

needles 

a fuzzy 

appearance. 

Courtesy of 

Sandy 

Gardosik, 

PDA 


All species of rust mites overwinter as eggs 

on the needles of host trees. Egg hatch 

occurs in very early spring, as early as mid- 

March. These are “cool-season” mites, like 

the spruce spider mite, but are active several 

weeks before spider mite eggs hatch. By the 

time spider mite overwintering eggs hatch, 

rust mites have completed several generations 

and rusty or bleached damage may be 

apparent (Fig. 8). 

During late spring and summer, rust mite 

populations are very low. However, in late 

summer when temperatures begin to moderate, 

populations build again. Overwintering 

eggs are deposited in fall, but rust mites may 

be active into December. 

White pine sheath mite biology may 

involve two distinctly different phases of 

activity. When candles are elongating, mites 

inhabit the needle sheath and feed on the 

elongating needles (Fig. 9). This causes 

needles to be stunted. The mites feed openly 

on mature needles, causing them to turn 

silver or brown. 


Strategies 



at this time. 

Preseason 

• Begin scouting in early March by examining 

10–20 trees per acre. Choose trees 

with previous damage or, if no previous 

damage has occurred, trees at random. 

— On spruce, look for clusters of red eggs 

on underside of needle. 

— On fi r, look for amber-colored eggs 

along the base of the needle where it 

lies fl at against the stem. 

— On pine, look at the exposed needle 

base of last year’s growth. 

• Scouting on overcast days will help 

symptoms stand out. 

• Look for predatory mites that may be feeding 

on overwintering eggs or immature 

mites following egg hatch. Predatory mites 

may keep this pest in check. 

Figure 8. Mite feeding damage 

giving foliage a rusted or 

bleached look. Courtesy of 


Figure 9. White pine sheath 

mites (circled). Courtesy of 


ERIOPHYID RUST AND SHEATH MITES ................................................................................................................................................................................................................................................. 43


• Growing degree days: Overwintering eggs 

hatch at 0–15 GDDs. 

• Threshold level (both criteria for rust 

mite on hemlock must be met): 

— At least 80 percent of sampled shoots 

have mites on them. 

— At least eight mites are present on 

one single needle in the plantation 

(examine all sides of the needle). 

• Continue scouting in spring until temperatures 

rise. Look for pine sheath mites by 

gently pulling elongating needles from the 

sheath and examining the needle bases. 

• Resume scouting in fall for population 

resurgence. 




Biological 

• Encourage predatory mites. 

Mechanical 


at this time. 

Biorational 

• Apply dormant oil in early spring before 

bud break. Eggs may have already hatched, 

but oil will control active mites if coverage 

is thorough. Oil will remove the blue color 

on some spruce. 

Chemical 

• Apply a miticide specifi cally labeled for 

rust or eriophyid mites in mid-April– 

mid-May. General miticides may not 

control eriophyid mites. Make a second 

application if warranted 1–2 weeks later. 

• Scout to determine if a fall application is 

necessary. 





Gypsy moths are of limited geographical 

distribution in the United States. They are 

responsible for signifi cant damage to many 

hardwood trees in the northeastern states. 

Regulations are in place for shipping cut 

trees from areas known to be infested with 

gypsy moth, including Pennsylvania. Contact 

the appropriate state or federal regulatory 

agency for information before shipping 

cut trees out of state. 

Hosts 

• Many hardwood species 

• Some conifers: pines, spruce, eastern red 

cedar, and occasionally fi rs and Douglas-fi r 


• Low–moderate 

• Heaviest damage frequently caused by 

more mature larvae that have consumed 

all the food supply in the area; repeated 

light or a single severe defoliation of 

conifers usually results in tree mortality 


• Defoliation of branches or entire tree 

(at fi rst noticeable at the top of the tree) 

• Tan egg masses on the trunk or underside 

of large branches 

• Gypsy moth caterpillars up to 2½ inches 

(63–64 mm) long 

• Reddish-brown pupal cases hanging from 

trunk 

• Adult moths in midsummer 


• None 


The oval, convex egg masses, up to 1½ 

inches (40 mm) long, are a mixture of hundreds 

of roundish, tan eggs and buff-colored 

hairs from the abdomen of the female. They 

can be found year-round attached to any 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

hard surface such as tree trunks or branches, 

fence rows, siding of homes, or anything left 

outside for the summer. Healthy egg masses 

are lighter in color; emerged egg masses are 

darker and have numerous holes in the surface. 

Holes may be from emerging caterpillars 

or parasitoids. 

Newly emerged gypsy moth caterpillars 

are dark brown, ¼ inch (6 mm) long, 

and have longer hairs at each end, which 

results in the larvae appearing somewhat “I” 

shaped. As the larvae mature, the characteristic 

colored spots on the upper portion 

of the body become apparent. Each of the 

fi rst fi ve segments of the larva’s body has 

a pair of blue spots; the next six segments 

each have a pair of red spots. Between these 

spots, a thin, yellowish line runs the length 

of the body. The head of a mature caterpillar 

is black and tan mottled, and numerous 

long, brown hairs cover the body. Caterpillars 

reach 2½ inches in length (63–64 mm) 

at maturity. 

The dark reddish-brown to black pupal 

cases are not surrounded by any protective 

silk cocoon. They are attached to the substrate 

by several strands of silk and the cast 

skin from the last instar caterpillar may be 

nearby. Female pupal cases are larger than 

male pupal cases. 

The gray-brown male moths emerge fi rst. 

They have a wingspan of about 1½ inches 

(40 mm) and are agile fl iers. They can be 

recognized by their rapid, zigzag fl ight as 

they search for female moths. This makes 

close examination diffi cult, but the males 

have prominent feathered antennae, and 

their wings have irregular dark lines across 

the forewings. The larger female moths 

have a wingspan of 2–2½ inches (50–63 

mm). Their wings are white with faint, dark 

markings. The heavy female’s body prevents 

fl ight, so they are usually found not far from 

their empty pupal cases. 


GYPSY MOTH 

Lymantria dispar 

(Linnaeus) 

Severe gypsy moth damage 

on Colorado blue spruce. 

Courtesy of PDA 

GYPSY MOTH ........................................................................................................................................................................................................................................................................................ ..................................................................... 29 45

Figure 1. Gypsy moth egg mass. 

Courtesy of Pennsylvania DCNR 

Forestry Archive, Bugwood.org 

(#5020037) 

Figure 2. Larvae “ballooning,” 

or dropping down onto new 

hosts via silken thread. 

Courtesy of Brian Schildt, PDA 

Figure 3. Gypsy moth larvae 

and silk covering Douglas-fi r 

buds. Courtesy of Brian Schildt, 

PDA 

Figure 4. Maturing caterpillar 

with red and blue spots and 

yellowish lines running the 

length of the body. Courtesy of 

Brian Schildt, PDA 


Eggs overwinter in the egg masses protected 

somewhat by the hairs from the female’s 

abdomen (Figs. 1 and 6). The masses may 

contain up to 600 eggs, which are rounded 

and tan. Egg masses darken with age, and 

hatching occurs between late April and 

mid-May when oak leaves are expanding. 

Newly hatched larvae (caterpillars) remain 

on the egg mass for a few days before moving 

to the newly developing leaves. If the 

population is high or the host is not suitable 

for development, larvae will “balloon” to 

new hosts by dropping down from a silken 

thread (Figs. 2 and 3). Spring winds can 

carry the young caterpillars up to a mile. 

During this phase, gypsy moth larvae are 

frequently seen on clothing when working 

outdoors. The irritating hairs of the fi rst 

instar caterpillars may cause contact 

dermatitis for some people. 

Larvae feed for 6–7 weeks, gradually 

increasing in size and appetite. Young larvae 

remain in the tree’s foliage day and night, 

but their feeding pattern changes when they 

are half grown (Fig. 4). The more mature 

larvae move down the tree to take refuge in 

a cool, shady place during the day and feed 

in the canopy at night. 

By late June to early July, the larvae fi nd 

a sheltered place to pupate (Fig. 5). About 

2 weeks are required before the adults begin 

to emerge, with the male moths appearing 

several days before the females. When 

the female moths emerge, they may crawl 

a short distance to an elevated spot or stay 

near the pupal case to emit a pheromone 

to attract the male moths. After mating occurs, 

females deposit one to two egg masses 

before dying (Fig. 6). Neither male nor 

female moths feed during their short lives. 

Egg masses overwinter until the following 

spring. One generation is produced per year 

in Pennsylvania. 

Figure 5. Gypsy moth pupae suspended from 

tree limbs. Courtesy of PDA 


Strategies 


• Plant the crop away from hardwood stands 

such as aspen or oak or other preferred 

hosts of the gypsy moth. 

Preseason 

• Scout year-round for gypsy moth egg 

masses both in the plantation and in 

surrounding woodlots. 

• Monitor egg masses for fi rst signs of 

hatching. 

• Contact a regulatory agency or county 

extension offi ce to obtain information 

about specifi c management options in the 

area and about shipping Christmas trees. 


• Growing degree days: 90–448 GDDs is 

the treatment window for the larvae. 

• When larvae are small, consider 

spraying with a microbial or other type 

of insecticide. 

• To help monitor populations, use a 

synthesized female pheromone trap to 

look for male moth emergence. 



Figure 6. Female 

gypsy moth over 

egg mass. Courtesy 

of Hannes Lemme, 

Bugwood.org 

(#1260007) 



Biological 

• Encourage natural predators of gypsy 

moths in all life cycle stages: egg parasitoid 

(Ooencyrtus kuvanae Howard), 

parasitoid of larva (Blepharipa pratensis 

Meigen and Meteorus pulchricornis 

Wesmael), predatory beetle (Calosoma 

sycophanta Linneaus), birds, mice, shrews, 

and ground beetles. 

• “Wilt” is a naturally occurring virus that 

is always present in the larvae but doesn’t 

kill them until they become stressed due 

to overcrowding or reduced food supply. 

• The fungal insect pathogen Entomophaga 

maimaiga can help decrease heavy populations 

during wet spring weather. 

Mechanical 

• In fall, winter, or spring, before eggs 

hatch, scrape egg masses off the trees 

or other areas where they are located. 

Destroy the eggs by burning or soaking 

them in soapy water for several days. 

Biorational 

• Use a microbial insecticide such as Bt 

(Bacillus thuringiensis) when larvae are less 

than 1 inch (25 mm) long for maximum 

control. 

Chemical 

• Several insecticides are registered for 

this pest. Read and follow all labels 

accordingly. 




GYPSY MOTH ......................................................................................................................................................................................................................................................................................... 47

LOPHODERMIUM 

NEEDLE CAST 

Lophodermium 

seditiosum Minter, 

Staley, and Millar 

Lophodermium needle 

cast, the “spring reddener,” 

infection on a fi eld of pine. 

Courtesy of Edward L. 

Barnard, Florida Department 

of Agriculture and Consumer 

Services, Bugwood.org 

(#4823057) 

Hosts 

• Scotch, Austrian, and red pines most 

susceptible 

• Most two- and three-needled pines 


• Moderate on fi eld-grown trees 

• High on seedlings 


• May fi rst develop in needles on lower 

branches 

Late Winter Through Early Spring 

• Infected needles develop yellow or 

reddish-brown spots 

• Current season’s infected needles redden 

by early spring 

Late Spring Through Midsummer 

• Needles brown and may fall from tree, 

leaving only new, healthy-appearing 

growth 

• Raised, black, football-shaped fruiting 

bodies appear on dead needles still 

attached to the tree and those on the 

ground 


• Winter burn 

• Dothistroma needle blight 

• Lethale needle cast 

• Cyclaneusma needle cast 



The most accurate and easiest identifi cation 

period for Lophodermium needle cast 

is in mid- to late summer when the fruiting 

bodies are present on needles infected the 

previous year. The shiny, black, footballshaped 

fruiting bodies are 1 ⁄32 inch (0.8 mm) 

long. They are slightly raised and aligned 

lengthwise on the needle. When mature, 

they have a longitudinal slit through which 

spores will be released. To see if fruiting 

bodies are mature, wrap some symptomatic 

needles in a moist paper towel for about 

20 minutes to see if they open and release 

spores. Fruiting bodies occur on dead 

needles remaining on the tree or those 

already on the ground. 

This common fungal disease is often referred 

to as the “spring reddener” of Scotch 

pines. In spring, needles infected the previous 

year will have small, brown spots, often 

with yellow margins. The most noticeable 

symptoms occur as the season progresses 

and needles turn yellow and eventually 

reddish brown. They may be cast anytime 

after symptoms appear. Fruiting bodies are 

required to confi rm a diagnosis. 


Needle infection starts in mid- to late summer 

and only the current year’s growth is 

susceptible. Spores released from mature 

fruiting bodies (Fig. 1) are dispersed by 

wind or water to the new growth or to other 

trees. The fungus enters the healthy needles 

through stomata and begins to disrupt the 

moisture-distribution mechanism in the 

needles. Symptoms are generally not apparent 

until early spring of the year following 

infection. At this time, infected needles begin 

to change color, fi rst yellowing and then 

turning reddish brown by the time shoots are 

elongating (Fig. 2). Severe infections will 

Figure 1. Lophodermium fruiting 

bodies. Courtesy of Tracey Olson, PDA 

Figure 2. Infected needles from the previous 

year turn reddish brown and drop in the spring. 

Courtesy of USDA Forest Service Northeastern 

Archive, Bugwood.org (#1398006) 


cause an entire tree to appear scorched, with 

only small, green shoots appearing at the 

end of branches. As the disease progresses, 

infected needles generally drop from the 

tree, leaving only the healthy, new growth. 

By late summer, fruiting bodies are mature 

and can be found on cast needles under 

the tree, lodged in branches, and on the few 

browned needles remaining on the branches 

(Fig. 3). When moisture is present, the 

fruiting bodies split longitudinally to release 

minute spores. This often occurs during cool 

rain events in late summer and fall. Since 

most of the infection comes from needles 

already on the ground, poor air circulation 

under the tree is a major contributor to the 

spread of the disease. The infection from 

needles already cast from the tree is unique 

to Lophodermium. Most other fungal diseases 

are spread from structures remaining on the 

tree. 

Figure 3. Characteristic football-shaped 

Lophodermium fruiting body. Courtesy 

of Tracey Olson, PDA 


Strategies 




nonshaded). 

• Do not plant near windbreaks of other 

susceptible conifers. 

• Remove infected trees around the block; 

rake and remove disease-bearing needles 

under infected trees. 

• Adequately space trees when planting to 

encourage drying and minimize disease 

transmission. 

• Avoid planting short-needled “Spanish” 

Scotch pine and “French Green” varieties 

since they are very susceptible to infection. 

• Examine all seedlings before planting 

and cull any suspected of being infected. 

Buy from a reputable producer. 

Preseason 

• Control weeds around and under trees to 

allow for good air circulation and drying 

of foliage. 

• In very early spring, scout for yellowing 

of needles. Needles often have small, 

brown spots with yellow margins that 

increase with time. Tag any suspect trees 

to examine later for reddening of needles. 

• Examine dead needles under suspect 

trees for evidence of fruiting bodies from 

previous year. 


• Scout for symptoms of Lophodermium in 

May and June by examining the lower 

branches of at least 50 trees of varying 

ages scattered throughout the block. Tag 

several suspect trees to watch for fruiting 

body production. 

• Threshold level: If at least 10 percent of 

the examined trees show injury, consider 

treating the entire plantation. 





Infection 

Symptoms 

Casting 


Bud Break 


LOPHODERMIUM NEEDLE CAST ............................................................................................................................................................................................................................................................ 49


Biological 


at this time. 

Mechanical 

• In spring, remove and destroy severely 

infected trees. Rake and remove any 

fallen needles and branches. 

• After harvest, examine needles around 

fresh stumps. If fruiting bodies are found, 

remove and destroy debris. 

• Avoid tip-ups (live branches on stumps) 

that may contain infected needles. 

Biorational 


at this time. 

Chemical 

• Apply an appropriate fungicide three to 

four times beginning in July and continuing 

at three-week intervals. If early 

summer is warm and/or wet weather is 

prolonged, adjust the timing and number 

of the sprays accordingly. 

• Apply the fungicide during periods of low 

air movement and when material will 

have adequate time to dry on needles. 


• Inspect plants/nursery stock and buy 


• Collect seeds from trees showing 

resistance for a seed-collection program. 


Hosts 

• Pines, especially Scotch and Mugo 

• Occasionally on Douglas-fi r, true fi rs, 

and spruce 


• High on Scotch pine 

• Low on other hosts 


Throughout the Year 

• White “fl ecks” on needles; white, 

oyster-shell-shaped scales that are 

generally heaviest on lower branches 

• Yellowing or brown needles leading to 

premature needle drop 

• Heavy infestations give a frosted or silvery 

appearance 

• Untreated infestations may result in death 

of twigs, branches, and eventually the tree 


• Needle cast diseases 


Pine needle scale is an armored scale that 

produces a white, oyster-shell-shaped, wax 

covering. This characteristic covering is 

easiest to fi nd on overcast days. The covering 

of the mature scale measures 1 ⁄16– 1 ⁄8 inch 

(2.5–3.0 mm) in length and has a small, 

yellow spot on the narrow end. Male scale 

insects are fragile, gnatlike, and capable 

of short fl ight but are rarely seen. Female 

pine needle scale insects are tan with 

legless, wingless, fl attened bodies beneath 

the white covering. Males do not feed. 

Females and immature scales have hairlike 

piercing mouthparts used to suck sap from 

needles. Pine needle scales do not produce 

honeydew as they feed. 

The red, oval eggs are found beneath 

the covering of the female. Newly hatched 

nymphs, or crawlers, are paprika colored. 

Crawlers must molt and settle to feed a 

short time after emerging from the egg. 

After they settle, the mouthpart is inserted 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

into the needle, preventing further movement. 

Settled nymphs are yellowish to tan. 

After a very short period, they begin to 

produce the characteristic white covering. 


Pine needle scale has two generations per 

year in Pennsylvania. Before dying in fall, 

each female deposits approximately 40 eggs 

(Fig. 1). These oval, red eggs overwinter 

under the dead female’s covering and begin 

hatch by mid- to late May. Egg hatch and 

subsequent crawler emergence may occur 

over a period of 2–3 weeks. The fl attened, 

red crawlers have legs but lack mouthparts 

(Fig. 2). They move around on the needle 

and must settle and molt in a short time. 

Crawlers may fall from needles or be blown 

by air currents to infest neighboring trees. 

They can also be carried by insects, animals 

(including people), and machinery. 

Figure 1. Overwintering pine needle 

scale eggs with dead female scale. 

Courtesy of Sandy Gardosik, PDA 

Figure 2. Pine needle scale crawlers. 



PINE NEEDLE 

SCALE 

Chionaspis pinifoliae 

(Fitch) 

White fl ecks on Scotch pine 

needles (signs of a pine 

needle scale infestation). 

Courtesy of Cathy Thomas, 

PDA 

PINE NEEDLE SCALE .............................................................................................................................................................................................................................................................................. ..................................................................... 29 51

Figure 3. Adult female pine 

needle scale. Courtesy of PDA 

Figure 4. Mature pine needle 

scale population. Courtesy of 


Figure 5. Twice-stabbed lady 

beetle feeding on pine needle 

scales. Courtesy of Rayanne D. 

Lehman, PDA 

Settled crawlers insert their long mouthparts 

into the needle and begin to secrete 

the armored covering. Initially, the covering 

is clear or yellow, but eventually the characteristic 

white covering is produced (Fig. 3). 

This generation feeds on the previous year’s 

growth and can readily be hidden by new 

growth in spring. 

By July, both winged males and wingless 

females are present. Following mating, egg 

laying, and hatching, the crawlers resulting 

from this generation move onto the current 

year’s needles and settle to feed. This generation 

is most noticeable (Fig. 4). By September, 

the second generation is complete and 

females begin to deposit overwintering eggs. 


Strategies 


• Properly space trees; allow enough space 

between saleable trees. 

• Remove and destroy any mature trees 

in and around the plantation acting as a 

source for infestation. 

Preseason 

• Scout for scales. 

— On an overcast day, look for scale 

coverings on lower branches. Pry off 

the scale covering with a pin and use 

a hand lens to look for overwintering 

eggs. If found, mark the tree to monitor 

for egg hatch. 

— Ragged chew holes or round holes in 

the scale covering indicate parasitoid 

or predator activity. 

• An application of dormant horticultural 

oil on infested trees may provide little to 

moderate success controlling overwintering 

eggs. Note: Oil will remove the blue 

color from blue spruces. 


• Growing degree days: 

— First-generation crawlers appear at 

298–448 GDDs. 

— Second-generation crawlers appear 

at 1,290–1,917 GDDs. 

• Threshold level: No specifi c threshold 

information is available at this time. 

Populations on Scotch and Mugo pines 

are capable of rapid increase, but those on 

other hosts are generally not signifi cant. 

• Treat if tree growth is stunted, yellowing 

occurs, or if scale populations become 

unsightly or aesthetically displeasing. 

Spot treat infested trees instead of treating 

the entire plantation. 

• Methods of scouting for crawlers (early 

May and early July): 

— Visit trees marked during late winter or 

early spring to look for crawler emergence. 

— Wrap electrical tape, sticky side out, 

around twigs with high populations 

of scales; monitor daily for the fi rst 

crawler to emerge. 

— Hold a white piece of paper under a 

branch while shaking the branch. 

Look for crawlers on the paper. 

• While scouting for crawlers, always scout 

for predators (several lady beetle species 

such as twice-stabbed lady beetle, 

lacewings, etc.) and wasp parasitoids or 

evidence of their presence. 




Biological 

• Encourage natural predators and 

parasitoids to assist with managing small 

infestations (Fig. 5). 

• If possible, reduce insecticide use for 

other pests. 

• Larger infestations may not be controlled 

with predators alone. 

Mechanical 

• Prune and destroy heavily infested 

branches; prune branches that are in 

danger of coming into contact with 

one another. 

• Do not mow or remove infested trees 

during crawler emergence as this may 

spread crawlers. 

Biorational 

• Insect growth regulators (IGRs) can be 

used during the crawler stage to disrupt 

the molting process. 

Chemical 

• Apply a spray targeting crawlers for 2–3 

weeks at 7-day intervals after the fi rst 

crawler is seen (most effective after egg 

hatch and before development of white 

wax covering). Note: This poses a risk 

to benefi cial predators and parasitoids. 

Whenever possible, use a “softer, 

predator-friendly” insecticide. 

• Systemic insecticides can be effective 

against young, settled nymphs that have 

just begun feeding (June and August). 





Hosts 

• European and redheaded pine sawfl ies: 

Scotch, red, Mugo, Jack, and Austrian 

pines 

• Introduced pine sawfl y: eastern white pine 


• Moderate 


• Complete defoliation or sparse, patchy, 

missing foliage anywhere on the tree 

• Needles may appear brown, wilted, and 

strawlike or twisted, as if singed 

• Clusters of young larvae feeding on 

needles 


• None 


Adult pine sawfl ies are seldom seen. They 

are related to and resemble bees in size and 

shape. They have two pairs of transparent 

wings but are not capable of stinging. 

Instead of a stinger, the female has a sawlike 

ovipositor that she uses to make a slit in the 

edge of a needle. She deposits a single egg 

into each slit and several eggs in a needle. 

The larvae are caterpillar-like with six 

or more pairs of prolegs on the abdomen. 

Moth and butterfl y caterpillars have fi ve or 

fewer prolegs. Both types of larvae also have 

three pairs of jointed true legs. Larvae use 

their chewing mouthparts to consume entire 

needles, which can result in extensive 

defoliation. 

Young larvae feed in colonies or clusters 

and can defoliate entire trees, depending on 

the size of the colony and tree. A distinguishing 

habit of the larvae is that they 

collectively rear back when a hand is waved 

over the cluster. 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 


Scout for European pine sawfl y eggs in needles. 

European Pine Sawfl y 

Larvae are dull gray green with a shiny 

black head (Fig. 1). They have light green 

and black stripes running the length of the 

body and are capable of growing to approximately 

1 inch (25 mm) long. Larvae feed on 

previous year’s growth, causing a decrease 

in growth rate but not tree death since they 

do not attack current growth. Young larvae 

eat only the outside of old growth, leaving 

the needles brown and strawlike behind 

the green, current growth. Older larvae eat 

the entire needle, leaving behind only the 

needle sheath. The adults are brown, fl ylike 

insects that are ½ inch (12 mm) long. 

Introduced Pine Sawfl y 

Larvae are black or dark brown with two 

dark stripes down the back and yellow and 

white patches on the sides. The head is 

black and shiny, and the underside is pale 

yellow or white (Fig. 2). They can grow up 

Figure 1. Colony of feeding European pine 

sawfl y larvae. Courtesy of Steven Katovich, 

USDA Forest Service, Bugwood.org (#5369986) 

Figure 2. Introduced pine sawfl y larva. 


PINE SAWFLIES 

European Pine Sawfl y, 

Neodiprion sertifer 

(Geoffroy) 

Introduced Pine Sawfl y, 

Diprion similis (Hartig) 

Redheaded Pine 

Sawfl y, Neodiprion 

lecontei (Fitch) 

Feeding damage from 

European pine sawfl y. 


PDA 

PINE SAWFLIES ...................................................................................................................................................................................................................................................................................... ..................................................................... 29 53

Figure 3. Colony of redheaded 

pine sawfl y larvae. Courtesy of 


Figure 4. European pine sawfl y 

eggs. Courtesy of Rayanne D. 

Lehman, PDA 

Figure 5. Colony of feeding 

European pine sawfl y larvae. 


PDA 

Figure 6. European pine sawfl y 

adult. Courtesy of Louis-Michel 

Nageleisen, Département de la 

Santé des Forêts, Bugwood.org 

(#2102003) 

to 1 inch (25 mm) long. The larvae feed in 

groups when young and singly as they mature. 

Depending on the generation, larvae 

may eat old needles or new growth. The 

cocoon of the introduced pine sawfl y is a 

strong, brown, semiglossy, textured cylinder. 

Adults are chunky and have black heads 

and thoraxes. Males are ¼ inch (7 mm) 

long with brown or black abdomens, and 

females are 1 ⁄3 inch (8 mm) long with black 

and yellow abdomens. 

Redheaded Pine Sawfl y 

Newly hatched larvae are white, unspotted, 

and have a brown or black head. As larvae 

feed and mature, they develop a red head 

and two to four rows of brown/black spots 

on the yellow body (Fig. 3). The last segment 

has a larger black patch on its sides. 

They can grow to 1 inch (25 mm) long. 

This species prefers younger trees in shaded 

areas. The larvae eat old needles fi rst, but 

heavy infestation or additional generations 

will cause new growth to be eaten as 

well. Young larvae eat the outside of the 

needles, leaving behind brown, strawlike 

needles, while older larvae consume the 

entire needle. The cocoon is a papery, yet 

tough, brown cylinder with rounded ends. 

Adult females are reddish brown and have a 

black abdomen with white spots. Males are 

slender and black with feathery antennae. 

Females tend to be larger than males. 


The three common pine sawfl ies in Pennsylvania 

have similar life cycles that differ 

in the overwintering stage, timing of egg 

hatch, and number of generations each year. 

Each species deposits eggs inside slits created 

by females in needles. Only fertilized 

eggs will result in females; unfertilized eggs 

produce males. When larvae are mature, 

they produce capsulelike cocoons in which 

they pupate. 

European Pine Sawfl y 

European pine sawfl ies overwinter as yellow 

eggs deposited in the needles (Fig. 4). The 

eggs hatch in April through mid-May. Larvae 

feed as a colony (10–100 larvae) and eat 

previous year’s growth through July (Fig. 5). 

Three to four larvae may be seen feeding on 

a single needle. When the larvae are fully 

grown, they drop to the ground and pupate 

around mid-August to early September. 

Adults emerge in mid- to late September 

and mate (Fig. 6). They only live a few 

days and do not feed. Using the sawlike 

ovipositor, females lay 6–8 eggs per needle 

and may use 10–12 needles for oviposition. 

One generation occurs per year. 

Introduced Pine Sawfl y 

Introduced pine sawfl ies overwinter as 

prepupae in cocoons on the ground in leaf 

litter. They pupate in early spring, with 

adults emerging in May through early June. 

Some prepupae may exhibit diapause, meaning 

they may overwinter additional seasons 

before becoming adults. Adults mate, and 

the females lay light bluish-colored eggs in 

the needles and cover them with a green, 

frothy substance. Females each lay approximately 

70 eggs with 10 eggs per needle 

(Fig. 7). Eggs hatch in roughly 2 weeks and 

colonies of larvae begin eating old-growth 

needles. As the larvae mature, they feed 

solitarily until fully mature (Fig. 8). Around 

early July, larvae spin cocoons and pupate a 

Figure 7. Introduced pine sawfl y eggs. 

Courtesy of John H. Ghent, USDA Forest 


Figure 8. Mature introduced pine sawfl y larva 

feeding solitarily. Courtesy of PDA 


short time afterward. Cocoons can be found 

among the needles, at the base of branches, 

and within bark crevices (Fig. 9). Adults 

emerge, peaking in early August. They mate 

and again lay eggs. These eggs hatch around 

one week later, and the larvae begin feeding 

on old- and current-growth needles. In 

September, the larvae drop to the ground 

to overwinter as prepupae. Two generations 

normally occur per year, although a partial 

or full third generation can occur if weather 

conditions are favorable. 

Redheaded Pine Sawfl y 

Similar to the introduced pine sawfl ies, the 

redheaded pine sawfl ies overwinter as prepupae 

and may exhibit diapause. They pupate 

in spring and adults emerge a few weeks 

later. Females deposit approximately 100 

eggs, which hatch around one month later. 

Larvae feed in colonies for 5–6 weeks. They 

begin feeding on old growth fi rst but will 

also consume new growth during a heavy 

infestation (Fig. 10). When mature, the 

larvae drop to the ground and spin a cocoon 

to overwinter. In some southern locations, a 

second generation is produced in the same 

season. In those cases, fi rst-generation larvae 

feed for a shorter time before pupating and 

creating the second generation. Populations 

of redheaded pine sawfl ies tend to be somewhat 

cyclic. Outbreaks of high populations 

resulting in heavy defoliation for several 

years are followed by periods of low populations 

with little damage. 


Strategies 


• Plant nonpine species that are not 

susceptible to sawfl y damage. 

Preseason 

• In winter and early spring, inspect trees 

for European pine sawfl y eggs deposited 

in the needles. Tag trees to monitor for 

larvae. 

• Remove trees that are larger than marketable 

size so they don’t serve as reservoirs 

for sawfl ies. 


• Encourage natural predators. 

• Scout for young larvae feeding on needles 

beginning in May. Look for strawlike, 

brown needles and missing foliage throughout 

the season. 

• Growing degree days: European pine 

sawfl ies emerge at 78–220 GDDs. 

• Threshold level: 

— For introduced pine sawfl ies and 

redheaded pine sawfl ies, treatment may 

be warranted if more than 50 random 

trees in a young plantation have 10 

larvae per tree or if they exhibit sawfl y 

damage. Treat older plantations when 

damage level will hurt sale. 

— For European pine sawfl ies, treatment 

may be warranted if there is a large infestation 

and the tree is within 3 years 

of harvest. 




Biological 

• Birds, rodents, parasites, viruses, and other 

predators can help decrease sawfl y populations 

but are often not enough to manage 

the pest in a plantation setting. 

Mechanical 

• Prune infested branches of trees not ready 

for harvest if plantation infestation is 

small. 

• Remove larvae by hand and squash or 

place them in soapy water for several days 

(small infestations only). 

Biorational 

• Apply a horticultural oil or insecticidal 

soap labeled for control of sawfl ies when 

larvae are very young. 

Chemical 

• Spot treat young larval infestations with 

a registered virus or insecticide labeled 

for sawfl y control. Note: Bt (Bacillus 

thuringiensis) will not control sawfl ies. 

• Apply an insecticide labeled for sawfl y 

control to entire plantation when young 

larvae are present if more than 25 percent 

of trees are infested. 

• Chemical treatment may be needed more 

than once per season, depending on the 

species of sawfl ies. 




Figure 9. Introduced pine 

sawfl y cocoon on twig. 

Courtesy of Rayanne D. 

Lehman, PDA 

Figure 10. Feeding damage 

from redheaded pine sawfl y 

larvae. Courtesy of Sandy 

Gardosik, PDA 

PINE SAWFLIES ....................................................................................................................................................................................................................................................................................... 55

PLOIODERMA 

NEEDLE CAST 

Ploioderma lethale 

(Dearn.) Darker 

Symptoms of needles 

infected with Ploioderma 

needle cast. Courtesy of 


Hosts 

• Austrian and red pines 

• Other two- or three-needled, hard-pine 

species 





branches 


• Infected needles develop reddish-brown 

spots 

• Spots may girdle needle and kill tips, 

leaving healthy, green base 

• Current season’s infected needles redden 

by early spring 

Late Spring Through Early Summer 

• In June, fruiting bodies may develop 

within spots and dead portion of needle; 

fruiting bodies appear as long ( 2 ⁄ 125– 1 ⁄ 5 

inch; 0.4–5.0 mm), black lines 

• Dead tips of infected needles with green 

bases may break off 

• Completely infected needles are cast 


• Other needle cast diseases, especially 

Lophodermium needle cast 


Ploioderma needle cast is a disease of hard 

pines. Although the infection occurs in 

spring, symptoms are not apparent until 

winter when yellow spots and bands appear, 

giving the needles a mottled appearance. 

During the following spring, the spots and 

bands girdle the needle, turning the tip 

straw brown or gray in color while the base 

of the needle remains green. Black, longitudinal 

fruiting bodies appear in the infected 

portion of the needles in June. Sporulation 

occurs after they mature, and the needles 

are cast to the ground by the end of summer. 

Ploioderma needle cast closely resembles 

Lophodermium needle cast in symptoms 

and disease cycle. However, Lophodermium 

fruiting bodies are elliptical to round, while 

those of Ploioderma are elongate, running 

longitudinally on the needle. 


During wet spring and early summer weather, 

colorless spores are released from fruiting 

bodies that have opened lengthwise on the 

needle (Fig. 1). Spores can be disseminated 

by wind or rain and will germinate on newly 

emerging needles close by. The fungus 

enters the needle through the stomata and 

the fi rst symptoms of infection are visible on 

the new growth by winter (Fig. 2). The next 

spring, fruiting bodies release spores and the 

process begins again. 

Figure 1. Infected needles with characteristic 

elongate, black fruiting bodies. Courtesy of 


Figure 2. Early infection of Ploioderma needle 

cast. Courtesy of Joseph O’Brien, USDA Forest 





Infection 

Symptoms 

Casting 


Bud Break 



Strategies 




nonshaded). 

• Plant resistant varieties suited for the 

environmental conditions. 

• Inspect nursery stock before buying and 

planting. 

• Adequately space trees to ensure air 

circulation and reduce moisture levels. 

Preseason 

• Maintain good weed control. 

• Scout needles in early spring for signs and 

symptoms of Ploioderma. 


• If Ploioderma is found in the plantation, 

consider treating with a fungicide in the 

spring during bud break. 




Biological 


at this time. 

Mechanical 

• Promptly remove and destroy branches 

and trees showing signs of Ploioderma. 

Biorational 


at this time. 

Chemical 

• To protect new growth in the spring, 

apply a fungicide when new growth 

is 2 inches long and needles are just 

beginning to emerge from the fascicles. 

• Two additional fungicide applications may 

be necessary at 14- to 21-day intervals to 

protect needles during the spore release 

and infection period. 


• Purchase and plant disease-free nursery 


PLOIODERMA NEEDLE CAST .................................................................................................................................................................................................................................................................. 57

RED-BAND 

(DOTHISTROMA) 

NEEDLE BLIGHT 

Mycosphaerella pini 

Rostr. 

Anamorph: 

Dothistroma septospora 

(Doroguine) Morelet 

Girdling lesions banding the 

needle. Courtesy of Tracey 

Olson, PDA 

Hosts 

• Austrian, Ponderosa, and Mugo pines 

highly susceptible 

• Scotch and red pines generally resistant 


• Moderate 


Late Summer Through Early Fall 

• Dark green bands on recently infected 

needles; bands may contain yellow or 

tan spots 

• Brown or reddish-brown bands and lesions 

on needles several weeks after infection 

Late Fall Through Winter 

• Brown, dead needle tips with the base of 

the needle remaining green 

• Dark brown or black, tiny fruiting bodies 

in the dead portion of the needle 


• Previously green base turns brown 

• Premature needle drop occurs (most 

severe on lower branches) 


• Brown spot needle blight 

• Lophodermium and Cyclaneusma needle 

casts 


• Winter injury 

• Environmental stress 


This disease, also referred to as Dothistroma 

needle blight, is called red-band needle 

blight because of the most common symptom 

found in the fi eld. Although initial 

symptoms include dark green bands on the 

needles, these are quickly replaced with 

brown or reddish-brown lesions. Only the 

base of the needle will be green, with the 

remaining portion tan or brown. Eventually, 

tiny, dark brown or black fruiting bodies are 

produced and release spores. The fruiting 

bodies are visible in the diseased area with a 

hand lens and are typically covered by a fl ap 

of needle epidermis. 


Red-band needle blight infection occurs 

throughout the growing season during 

wet periods. Beginning in May, spores are 

released from dark brown or black fruiting 

bodies that rupture through the epidermis 

of previously infected needles (Fig. 1). The 

spores are spread by wind and splashing 

rain and enter healthy needles through 

the stomata. Second-year or older needles 

are susceptible to infection throughout 

the season. Current-year needles are not 

susceptible until they have elongated and 

hardened off, usually around July. Infection 

from cast needles on the ground below the 

tree is negligible. 

Figure 1. Fruiting body rupturing the 

epidermis. Courtesy of Tracey Olson, PDA 

After infection takes place, symptoms 

are not visible for 3–6 months on average. 

In late summer, infected needles may show 

dark green bands that look water soaked. 

This symptom occurs briefl y and is usually 

not detected. Next, yellow to tan spots become 

visible and then eventually change to 

become larger, reddish-brown lesions banding 

the needle (Fig. 2). The fungus produces 

a toxin that quickly kills the tissue at the 

infection site and causes it to turn purplish 

red, giving this disease its common name, 

red-band needle blight. Girdling lesions 

will result in needle tips dying while the 

bases of the needles remain green (Fig. 3). 

By late fall, tiny, black fruiting bodies may 

appear in the bands or dead tissue. These 

will not mature and release spores until the 

following spring. 




Infection 

Symptoms 

Casting 


Bud Break 


Figure 2. Reddish-brown lesions banding 

the needle. Courtesy of Tracey Olson, PDA 

Figure 3. Dead needle tips resulting from 

girdling by lesions. Courtesy of Tracey 

Olson, PDA 

Needle casting begins in fall and continues 

through late summer of the following 

year. The fi rst to drop are the second-year 

and older needles that were infected in 

spring (Fig. 4). Needles infected in the same 

year they are produced are usually not cast 

until late summer of the following year. 


Strategies 


• Plant resistant or tolerant varieties such as 

Scotch or red pine. 

• Plant in an area with good drainage and 

airfl ow. 

• Plant trees with adequate spacing to allow 

for airfl ow. 

Preseason 

• Control weeds in and around trees. 

• Remove and destroy lower whorls of 

branches to increase air circulation. 

• Scout for yellow needle spots, dead needle 

tips, and fruiting bodies in late fall or early 

spring; use a 15X hand lens to see the 

fruiting bodies. 


• If symptoms of disease are present, consider 

treating the plantation. Severely 

infected trees provide a good source for 

inoculum and should be removed. 

• Do not shear trees in wet weather. 

Shearing trees when foliage is wet may 

further spread the disease. 

• If disease is present in a block, shear 

disease-free stock fi rst. 

• Maintain weed control practices. 




Biological 


at this time. 

Mechanical 


at this time. 

Biorational 


at this time. 

Chemical 

• Apply an appropriate protective fungicide 

once in mid-May to protect older foliage 

and again in mid-July to protect needles 

of all ages. 




Figure 4. Infected previous 

years’ needles will soon be 

cast. Courtesy of USDA Forest 

Service Archive, Bugwood.org 

(#2251050) 

RED-BAND (DOTHISTROMA) NEEDLE BLIGHT ...................................................................................................................................................................................................................................... 59

RHABDOCLINE 

NEEDLE CAST 

Rhabdocline weirii 

A. K. Parker and J. Reid 

Rhabdocline 

pseudotsugae Syd. 

Foliage with Rhabdocline 

needle cast infection. 

Courtesy of Tracey Olson, 

Host 






branches 

Late Summer 

• Yellow spots or fl ecks on current-year 

needles that enlarge with time 


(Before Bud Break) 

• Reddish-brown spots on upper surface 

of current-year needles; distinct border 

between diseased area and healthy, green 

tissue 

• Swollen, light tan fruiting bodies on the 

underside of symptomatic needles 

Bud Break 

• Fruiting bodies rupture underside of 

needle epidermis, releasing mass of 

orange spores 

Late Spring (After Bud Break) 

• Darkened fruiting bodies indicating 

sporulation has passed 

Summer Through Early Fall 

• Previous year’s infected needles drop or 

cast; severely diseased trees might only 

retain current-year needles 


• Cooley spruce gall adelgid 

• Swiss needle cast 

• Douglas-fi r needle midge 


Douglas-fi r is the only known host of 

Rhabdocline. Laboratory analysis provides 

the most accurate identifi cation, but growers 

can generally identify this fungus in their 

fi elds by observing symptoms prior to bud 

break. In late winter or very early spring, 

look for reddish-brown splotches on the 

upper needle surface (Fig. 1). There is a 

distinct division between diseased tissue 

and the surrounding healthy, green tissue. 

As bud break approaches, remove shoots 

containing suspect needles and place them 

in a glass of warm water for a few minutes. 

Mature fruiting bodies of Rhabdocline will 

rupture the epidermis longitudinally on the 

underside of the needles and release a mass 

of orange spores. 


Rhabdocline needle cast infection occurs 

around bud break, when buds are opening to 

expose susceptible immature needles (Fig. 2). 

During periods of high humidity (rain shower 

or heavy morning dew), the mature fruiting 

bodies open (Fig. 3). Fruiting bodies are only 

found on the previous year’s needles and 

open on the underside of the needle. During 

spore release, the needle epidermis splits 

lengthwise, revealing a light tan to orange 

spore mass. Individual spores are dispersed 

by water and wind to the susceptible new 

growth. Spores germinate on the wet needle 

and penetrate the cuticle before the fungus 

begins to grow inside the needle. 

PDA Figure 1. Distinct symptomatic 

reddish-brown splotches of 

Rhabdocline needle cast (March). 


Figure 2. Early bud break of 

Douglas-fi r buds signaling the 

start of Rhabdocline sporulation. 


Figure 3. Fruiting bodies rupture 

on the undersides of Rhabdoclineinfected 

needles during bud break. 





Infection 

Symptoms 

Casting 


Bud Break 


The infection period may last several 

weeks, but only the newly emerging spring 

growth can become infected. When the 

fruiting body turns dark brown or black, 

spore production is complete for the year 

(Fig. 4.) Needles containing spent fruiting 

bodies will be cast from the tree through 

late spring and summer. Trees that have 

severe infection over consecutive years may 

hold only the most current year’s needles 

during the winter (Fig. 5). 

Figure 4. Brown to black fruiting 

bodies signaling that spore production 

is complete for the year. 


Figure 5. Severely infected tree with only 

current-year needles remaining. Courtesy of 

Brian Schildt, PDA 


Strategies 


• Choose a site with slope, good air circulation, 

and water drainage to promote 

drying of trees. 

• Properly space trees to encourage 

adequate drying of needles, especially 

in spring. 

• Plant resistant or tolerant tree varieties; 

avoid Douglas-fi rs from Rocky Mountain 

seed sources. 

• Remove and destroy unmanaged Douglasfi 

r that may act as a source of inoculum. 

Preseason 

• Maintain proper weed control year-round. 

• Concentrate scouting on trees planted in 

areas that are more conducive for disease 

development (low-lying or shaded areas). 

Look at all sides of the tree. 

• Scout the block in late winter or early 

spring (before bud break) for reddishbrown 

blotches on upper needle surface of 

most current-year needles. Overcast days 

afford the best opportunity to see needle 

discoloration. Pay particular attention to 

needles on the lower third of the tree and 

trees that look thin and only have last 

year’s needles. If symptomatic trees are 

located, tag several to observe for maturation 

of fruiting bodies. 

• Soak suspect needles in warm water to 

check for the maturity level of the fruiting 

bodies. When fruiting bodies open, spores 

are mature and ready to be released. 


• Threshold level: Ask a state/regional plant 

inspector about any regulatory thresholds. 

• In late April/early May, begin regular 

scouting for bud break in each target 

block. Make the fi rst fungicide application 

when the fi rst buds begin to open. 



RHABDOCLINE NEEDLE CAST ................................................................................................................................................................................................................................................................ 61


Biological 


at this time. 

Mechanical 

• Remove and destroy severely infected 

trees prior to bud break. 

• Prune and remove dead/dying branches 

when they are dry. 

• Butt-prune large trees to encourage drying 

of lower branches. 

Biorational 


at this time. 

Chemical 

• Apply an appropriate fungicide when buds 

begin to break on the trees in the block. 

Do not wait for new growth to elongate. 

Successful prevention of infection depends 

on preventing spores from infecting new 

needles—keep a protective coating of 

fungicide on the new growth for this to 

happen. 

• Follow through with two to three additional 

applications until needles are fully 

elongated and mature or fruiting bodies 

have darkened. See the schedule below: 

— First application: when fi rst trees in 

plantation break bud 

— Second application: 1 week after fi rst 

application 

— Third application: 2 weeks after second 

application 

— Fourth application: 3 weeks after third 

application only if spring is prolonged 

by cool, wet weather and sporulation 

remains active or Swiss needle cast is 

detected 





Hosts 

• Colorado blue spruce and Engelmann 

spruce most susceptible 

• White spruce occasionally susceptible 


• Moderate 



branches 

• Older needles may exhibit discoloration 

throughout the year 

Late Fall Through Early Spring 

• Yellow mottling of fi rst-year needles 

Early Spring (Before Bud Break) 

• Yellow needles turn brown or purplebrown; 

black fruiting bodies with white 

cap protruding from needle stomata 

develop 

Late Spring (After Bud Break) 

• Initial casting of previous year’s infected 

needles 

Summer Through Early Fall 

• Casting of previous year’s infected 

needles, leading to large bare areas on 

trees; severely diseased trees might only 

retain current-year needles 



• Cytospora canker 

• Flyspeck 

• Environmental stress (drought) 

• Nutritional defi ciencies 

• Spray damage 


Rhizosphaera needle cast disease is caused 

by the fungus Rhizosphaera kalkhoffi i Bubak. 

Colorado and Engelmann spruce are very 

susceptible, but Norway and white spruce 

are somewhat resistant. Young trees sustain 

the most severe damage, but trees of any 

size can be affected. Field identifi cation is 

based on symptoms, but accurate identifi cation 

requires laboratory analysis. 

Healthy spruce trees will retain 5 or 

more years of needles. Those infected with 

Rhizosphaera may have only one year of 

needles. Foliage on inner branches and on 

the lower portion of the tree is more prone 

to infection since these areas tend to remain 

wet for longer periods. When the infection 

is severe, this disease will kill lower branches 

and infection will progress to the upper 

branches. 

Yellowing needles of any age or any that 

are reddish brown to purple in fall are suspect. 

Look particularly at trees that appear 

thin on the lower half or are only retaining 

1–2 years of needle growth. Examine apparently 

healthy needles on these trees in early 

spring, prior to bud break. Look for minute, 

black fruiting bodies, 1 ⁄250 inch (0.1 mm) in 

diameter, in neat lines running the length of 

the needle. 


Rhizosphaera overwinters in infected needles 

on the tree. In spring, the fruiting bodies 

mature and push through the stomata. They 

appear as neat lines of tiny, black spots 

about the size of the stomata (Fig. 1). About 

the time of bud break, they mature, and 

when there is adequate moisture, spores are 

released. The spores are splashed by rain to 

the new growth, where infection will begin. 

The spores enter the stomata of the healthy 

needle and germinate after a minimum of 

48 hours. Under less favorable conditions, 

longer germination time is required. Most 

infection occurs in spring, but fi rst-year 

needles are susceptible throughout the 

growing season. In some areas of Pennsylvania, 

a second infection period in August and 

September has been reported. 

The disease continues to develop in 

the needle for the next year, but symptoms 

are generally not well expressed. This long 

period between infection and appearance 

of symptoms makes this disease diffi cult to 

diagnose and control. Around May, one 

year following infection, apparently healthy 

Figure 1. Fruiting bodies pushing up through 

the stomata. Courtesy of Tracey Olson, PDA 

RHIZOSPHAERA 

NEEDLE CAST 

Rhizosphaera kalkhoffi i 

Bubak 

Rhizosphaera-infected 

foliage on inner branches 

and lower parts of the tree. 

Courtesy of USDA Forest 

Service North Central 

Research Station Archive, 


RHIZOSPHAERA NEEDLE CAST ............................................................................................................................................................................................................................................................. ..................................................................... 29 63



Infection 

Symptoms 

Casting 


Bud Break 


Figure 2. Apparently healthy 

needles bearing fruiting bodies. 


Figure 3. Infected reddishbrown 

needle in fall. Courtesy 


Figure 4. Striking symptomatic 

color on Colorado blue spruce. 


Service Archive, Bugwood.org 

(#2634062) 

needles will bear mature fruiting bodies 

(Fig. 2). During summer, these now previous 

year’s needles will turn yellow and gradually 

reddish brown or purple by fall (Fig. 3). 

This discoloration is very striking, particularly 

on Colorado spruce with bright blue 

color (Fig. 4). Most infected needles are cast 

in the fall, but some infected needles will 

remain on the tree. 


Strategies 


• Plant on a slope with good drainage. 

• Plant resistant varieties such as Norway 

or white spruce. Plant disease-free stock. 

• Adequately space trees when planting 

to allow for air circulation and drying 

of needles. 

Preseason 

• Maintain proper weed control throughout 

season by mowing grass and brush. 

• Scout for fruiting bodies in spring before 

bud break. 

— Randomly select at least 20 trees to 

sample. Concentrate scouting on 

trees planted in areas that are more 

conducive for disease development. 

— Using a 10X hand lens, examine the 

2-year-old needles found on three 

lower branches of each tree sampled. 

Look for black fruiting bodies 

protruding from stomata openings. 

— During dry weather, remove and 

destroy infected branches and trees. 


• If at least half of the sampled branches 

have fruiting bodies on 10 percent of 

needles, consider treating the entire 

plantation with a fungicide application. 

• Shear healthy trees fi rst during dry 

conditions. 




Biological 


at this time. 

Mechanical 

• Cut infected branches back to the main 

trunk. 

• Do not leave live branches on the stumps 

of harvested trees. 

Biorational 


at this time. 

Chemical 

• Apply an appropriate fungicide when 

new shoots are ¾–1¼ inches long (needles 

are half elongated) and a second spray 

3 weeks later (needles are fully elongated). 

Moderately infected trees may require 

2 years of fungicide applications. 

• In some areas, it may be necessary to make 

another application in mid-August to 

early September to prevent late summer 

infection. 


• Maintaining tree health and vigor can 

help guard against severe Rhizosphaera 

needle cast infection. 


Hosts 

• Most spruce species susceptible 

• In Pennsylvania, most commonly reported 

on Colorado blue spruce and its varieties, 

Serbian spruce, and Sitka spruce 




Midsummer 

• Pale yellow bands become visible on 

current-year needles 

• Needles infected in previous year turn 

brown and defoliate 

Late Summer 

• Pale yellow bands with orange fruiting 

bodies (called telia) obvious on most 

recent growth; band will go entirely 

around needle and increase in color 

intensity as season progresses; discolored 

bands easiest to see on overcast days 

Bud Break 

• Bright orange telia mature and rupture on 

infected needles 


• Infected trees may appear disfi gured and 

have extensive needle discoloration, 

reduced growth, and premature needle 

drop 


• Mechanical damage from needle 

punctured by needle tips 

• Cinara aphids 

• Pesticide phototoxicity 


Weir’s cushion rust (Chrysomyxa weirii) is the 

only species of spruce needle rust currently 

found in Pennsylvania. Infected current-year 

needles will exhibit pale yellow bands that 

go completely around the needle. Orange, 

immature fruiting structures may be visible 

within the band. This is easiest to see on 

an overcast day or on the shady side of the 

tree. Mechanical damage from pricking of 

needles during windy weather may mimic 

spruce needle rust. But, mechanical damage 

does not encircle the needle and usually has 

a darkened, wrinkled spot or indentation in 

the yellowed area. Around bud break, the 

mature, orange telia form on the discolored 

spots. Spores are yellow orange. 

Another species of spruce needle rust, 

Chrysomyxa ledi, has been detected in surrounding 

states. The fruiting body of this 

fungus resembles white blisters on current 

growth and requires an alternate host. 


This species of spruce needle rust does not 

have an alternate host. All necessary spore 

stages are found on spruce. During the winter, 

the fungus is maturing inside the current 

year’s needles and faint yellow bands with 

developing orange telia can easily be seen, 

particularly on lower branches (Figs. 1 and 

2). As bud break approaches, telia become 

swollen and contrast with the green or blue 

of the needle (Fig. 3). 

Figure 1. Early symptoms of 

developing rust on Serbian spruce. 


Figure 2. Faint yellow and orange 

bands with maturing fungus inside 

needles. Courtesy of Tracey Olson, 

PDA 

Figure 3. Swollen telia, or fruiting 

bodies (April). Courtesy of Tracey 

Olson, PDA 

SPRUCE 

NEEDLE RUST 

Chrysomyxa weirii 

(H. S. Jacks) 

Yellow bands containing 

faint orange fruiting bodies 

distinctive of spruce needle 

rust. Courtesy of Tracey 

Olson, PDA 

SPRUCE NEEDLE RUST ........................................................................................................................................................................................................................................................................... ..................................................................... 29 65



Infection 

Symptoms 

Casting 


Bud Break 


Figure 4. Spores being released 

from the ruptured needle surface. 


PDA 

Figure 5. Sporulation occurs 

during bud break. Courtesy 


At bud break the blisters burst, releasing 

massive amounts of yellow-orange spores 

(Fig. 4). Wind and splashing rain carry 

spores to newly emerging needles on the 

same tree or adjacent trees. The infection 

period can continue for 2–3 weeks after 

bud break, and infection rate increases with 

high moisture. 

The disease cycle starts again with the 

newly infected needles harboring the disease 

until bud break the following year. Previously 

infected needles that have already 

dispersed spores will turn rusty brown (Fig. 

5), die, and drop from the tree in summer. 

Spores will not continue to be produced on 

dead needles. 


Strategies 


• Thoroughly inspect all seedlings before 

planting. 

• Space trees to encourage drying. 

• Control weeds to reduce the moisture 

level around the trees and to increase 

coverage if control measures are needed. 

• Remove and destroy abandoned hosts 

around fi eld to eliminate sources of 

inoculum. 

Preseason 

• Scout trees in winter for yellow bands 

with orange spots in the center indicating 

infection. Scout on overcast day or look at 

shaded needles to make it easier to detect 

discoloration. Lower branches are often 

fi rst to be infected. 

• If banding of needles is found, tag several 

trees to monitor for telia production. 

• Scout trees for orange telia beginning 1–2 

weeks before bud break and continue until 

2–3 weeks after bud break. 

• If only a few trees are found with symptoms, 

cut and remove them from the fi eld well 

before fruiting bodies mature at bud break. 


• Threshold level: No threshold level has 

been established for this fungus. If the 

fungus is present, it can spread rapidly 

with favorable environmental conditions. 

• This can be a more serious concern for 

Christmas tree growers who also dig trees 

since it is a disease of regulatory concern 

for many states. Ask state/regional plant 

inspectors about a regulatory threshold for 

this disease. 

• Apply a fungicide spray when trees have 

broken bud; make subsequent applications 

at weekly intervals until needles are 

mature or the symptomatic needles have 

dropped to the ground (approximately 

three to fi ve sprays total). 




Biological 


at this time. 

Mechanical 

• Remove and destroy infected trees before 

spore production and release. 

Biorational 


at this time. 

Chemical 

• Apply a fungicide spray when trees have 

broken bud; make subsequent applications 

at weekly intervals until needles are 

mature or the symptomatic needles have 

dropped to the ground (approximately 

three to fi ve sprays total). 


• Do not purchase or accept infected 

nursery stock. 


Hosts 

• All conifers, especially spruce and Fraser, 

Canaan, and balsam fi rs 


• High 


• Small, irregularly shaped yellow spots 

(“stippling”) on needles; on fi r, needle 

base may be pale 

• Rusty or bronzed needles; damage may appear 

most severe during hot, dry weather 


• Damage heaviest at the bottom inside of 

the tree; damaged needles will not recover 

from the chlorophyll lost as a result of 

mite feeding 

• Fine webbing on needles and twigs; cast 

skins, dead mites, dirt, and other debris 

trapped in the silk 

• Infestations frequently occur in pockets, 

not distributed evenly through fi eld 


• Rust mites (spruce and fi r) 

• Rhizosphaera needle cast (spruce) 

• Air pollution 

• Aphids (spruce) 


A 15–20X hand lens or small microscope 

is required to view spider mites and eggs. 

Adult spruce spider mites are oval and only 

1 ⁄50 inch (0.5 mm) long. They have thin 

hairs, or setae, on the top of their convex 

bodies. The body color depends on the host 

and varies from pale green to dark green 

or dark red; legs are generally pale in color. 

The fi rst stage, a six-legged larva, is salmon 

colored until it has fed for a short time. 

After feeding, the larva and all subsequent 

stages are generally green or dark red. The 

eggs are rounded and vary from tan (active 

season eggs) to red (overwintering eggs). 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

Each egg has a single hairlike stripe on the 

top, which can be used to distinguish spruce 

spider mite eggs from other spider mite eggs 

that may be found on conifers. 

Two other spider mites occur on Christmas 

trees: two-spotted spider mite and 

admes spider mite. The two-spotted spider 

mite is generally green with two dark spots, 

one on either side of the oval body. It is 

slightly smaller than spruce spider mite and 

overwinters as an orange female. The admes 

spider mite has only been found on spruce 

and is larger than spruce spider mite. The 

body is more fl attened and reddish brown 

with long, pale legs. The admes mite overwinters 

as an egg. 


Spruce spider mites can develop from an egg 

to an adult in 2–3 weeks under normal conditions 

(Figs. 1 and 2). It is a cool-season mite 

and attains the highest populations in spring 

and fall; temperatures above 80–90°F will 

result in population decline. During the hot, 

dry weather in summer, spruce spider mites 

seem to disappear, only to reappear when 

temperatures moderate in late summer or fall. 

Spider mites are still present during summer, 

but feeding and reproduction are greatly 

reduced. Predatory mites are very common 

on the host plants at this time of year. 


Monitor for egg hatch. 

Apply dormant oil. 

Figure 1. Adult 

spruce spider 

mite. Courtesy 

of Rayanne D. 

Lehman, PDA 

Figure 2. Adult 

spruce spider 

mite. Courtesy 

of Rayanne D. 

Lehman, PDA 

SPRUCE 

SPIDER MITE 

Oligonychus ununguis 

(Jacobi) 

Spruce spider mite feeding 

damage. Courtesy of 

Eric R. Day, Virginia Tech, 


SPRUCE SPIDER MITE ............................................................................................................................................................................................................................................................................ ..................................................................... 29 67

Figure 3. Overwintering spruce 

spider mite eggs. Courtesy of 


Figure 4. Newly molted spruce 

spider mite adults. Courtesy of 


Figure 5. Chlorotic spots resulting 

from spider mite feeding. 


Figure 6. Spider mite feeding 

damage on older growth. 

Courtesy of USDA Forest Service 

Northeastern Area Archive, 


Spruce spider mites follow the typical 

spider mite life cycle: egg, six-legged larva, 

two eight-legged nymphal stages, and adult 

(male or female) (Figs. 3 and 4). Between 

each of the active stages, a resting stage, 

or chrysalis, is formed as the mite prepares 

to shed its outer skin. This molting stage is 

generally not affected by chemical controls. 

The male completes development fi rst and 

may be seen “guarding” the female chrysalis 

until she emerges and mating occurs. Mated 

females produce both male and female offspring, 

while unmated females produce only 

females. Depending on temperature, each 

female can deposit an average of 30–40 eggs 

during her lifetime. After the fi rst generation, 

overlap of generations is common and 

all stages are present at any given time. 

Spider mites have piercing-sucking 

mouthparts. As they feed, they withdraw 

sap containing chlorophyll from the needles 

(Fig. 5). They prefer to feed on older growth 

and will generally not feed on current-year 

needles until they have hardened off (Fig. 

6). This species generally does not produce 

copious amounts of silk. But, when populations 

are high, larvae can spin down on the 

silk threads they produce and be carried by 

air currents, insects, birds, or other animals 

to new host plants (Fig. 7). Six or more 

generations occur a year in Pennsylvania. 

Figure 7. Spider mite webbing buildup 

during a more severe infestation. Courtesy 

of USDA Forest Service Region 4 Archive, 



Strategies 


• During site selection, consider elevation 

and aspect of the fi eld. Warmer temperatures 

generally associated with southfacing 

slopes and lower elevations favor 

spruce spider mite populations. 

• Drought-prone locations favor population 

buildup. 

• Inspect nursery stock for infestations. 

Preseason 

• Manage groundcover and surrounding 

area to encourage natural predators. 

• Growing degree days: Based on observations 

in Pennsylvania, egg hatch occurs at 

50–121 GDDs. 

• Scout trees for overwintering eggs in 

very early spring with a 15X hand lens. 

Examine shoots for red, rounded eggs usually 

located on twigs at the base of needles 

or around buds. Monitor for egg hatch 

by tagging one or more trees known to 

have a population of overwintering eggs. 

Hatch will generally begin on the south 

side of the tree. Make observations every 

few days to determine when eggs begin 

to hatch. Egg hatch occurs prior to bud 

break. Record percentage of egg hatch if 

miticide use is planned. 

• Scout for active forms by holding a 

light-colored surface (paper, paper plate, 

painted clipboard, inside back cover of 

this manual, etc.) beneath a branch. Tap 

the branch sharply by hand or with a stick 

(Fig. 8). Mites will be dislodged and drop 

to the monitoring surface. Tilt the surface 

slightly to allow dirt and debris to slide off 

and then observe for mites. Spider mites 

will begin to crawl and appear as minute 

dark dots moving slowly over the surface. 

Examine with your hand lens to verify 

that they are spider mites. Sample several 

branches around the tree and multiple 

trees per block to determine population 

level. 

• Smearing suspected mites can be another 

test—dark green or dark red smears on the 

monitoring surface usually indicate spruce 

spider mites. 

• Populations can be estimated by examining 

shoots of current growth taken from 

the bottom third of trees. Use a hand lens 

to check for active forms or viable eggs. 

Sample about 15 shoots per acre or a minimum 

of 10 shoots in a small block; sample 

every part of block as populations tend to 


e spotty in a fi eld. Determine the percentage 

of shoots that have mites or mite 

eggs. Repeat this procedure every several 

weeks to detect population fl uctuations. 

• Emerald green arborvitae is a good indicator 

plant and will show signs of infestation 

early. If planted, monitor closely for fi rst 

signs of spruce spider mites. 


• Continue monitoring populations using 

the beating method or by observing eggs 

or active mites on branches. Repeat 

sampling every 3–4 weeks throughout the 

growing season and within one week following 

pesticide application. 

• Watch for increase in population numbers, 

which may require control. Damage 

observations, although helpful in locating 

populations, are not accurate measures of 

population levels. 

• Drought and/or warm weather around 

autumn could delay fall egg hatch. 

• Growing degree days: Peak mite activity 

occurs at 192–363 and 2,375–2,806 

GDDs. 

• Threshold level—two methods described 

below determine the economic threshold: 

— Mites per branch: More than 10 mites 

per branch on most branches sampled 

indicate that pesticide intervention is 

recommended. 

— Percentage of infested trees: Divide 

the number of shoots with mites and/or 

mite eggs by the total number of shoots 

examined and multiply by 100 to 

calculate the percentage. Refer to the 

table below to determine if pesticide 

intervention is recommended. 

THRESHOLD LEVEL 

Economic 

Size of Tree 

Threshold* 

Less than waist high 

(< 3.2 feet) Up to 40% 

Waist high (3.2 feet), 

year before sale Up to 20% 

Greater than waist 

high (> 3.2 feet), 

year of sale Up to 10% 

*Threshold levels may be different for trees 

that will be dug as opposed to cut. Check 

with your state/regional plant inspector for 

acceptable mite levels. 

• At the end of the season, update records 

and evaluate results. 


Biological 

• While scouting for spruce spider mites, 

also look for natural predators such as 

fast-moving, pale phytoseiid mites, hover 

fl y larvae, lacewing larvae, dusty wings, 

and lady beetles. All are effective predators 

and may help control the population 

of spruce spider mites. Refer to Appendix 

B: Biological Controls Photo Chart for 

pictures. 

Mechanical 

• Streams of water under pressure can effectively 

wash mites from trees. However, this 

is not practical in the fi eld situation, but 

long periods of heavy rain can signifi cantly 

reduce mite populations. 

Biorational 

• Dormant oil applied prior to egg hatch 

can reduce the population. Note: Oil will 

remove the blue coloring from spruce. 

Chemical 

• General insecticides are not as effective 

for control as are specifi c chemicals referred 

to as miticides. General insecticides 

are often detrimental to populations of 

benefi cial insects and mites. 

• Numerous miticides are available for controlling 

spruce spider mite on Christmas 

trees. Ovicides that target eggs and materials 

that control both eggs and active forms 

often provide the longest control. Ovicides 

generally do not work on overwintering 

eggs. For best results, apply controls when 

population is low or starting to build. 

Evaluate control after 5 days. 

• A second application may be needed in 

7–10 days, unless prohibited on the label. 

Some labels recommend application one 

time per year or even every other year to 

lessen chance of resistance developing in 

the population. 

• Spider mites are capable of quickly becoming 

resistant to a particular miticide 

due to their rapid reproduction. To avoid 

resistance, alternate classes of chemicals 

every third application. 




Figure 8. Tapping branches over 

white paper to dislodge mites. 


SPRUCE SPIDER MITE ............................................................................................................................................................................................................................................................................. 69

SWISS 

NEEDLE CAST 

Phaeocryptopus 

gäumannii 

(T. Rohde) Petr. 

Foliage with symptoms and 

signs of Swiss needle cast 

infection. Courtesy of Tracey 

Olson, PDA 

Host 





• May fi rst develop on lower branches 

Spring Through Fall 

• Two parallel rows of tiny, black fruiting 

bodies (pseudothecia) on the underside 

of fi rst- , second- , or third-year needles 

• Fruiting bodies may not develop on 

current-year infected needles until after 

a frost 

• Dieback of needle tips resembling drought 

damage 

• Yellowing or mottling of previously 

infected needles 

• Browning and casting of needles infected 

at least one year prior; heaviest casting 

seen on interior of lower branches 



• Flyspeck 

• Cooley spruce gall adelgid 

• Environmental stresses 

• Nutrient imbalances 

• Winter burn 


Symptoms are present on needles within 

3 years of infection. The fruiting bodies of 

Swiss needle cast are easily detected and 

can be seen with a hand lens anytime 

throughout the year. Look for two bands of 

tiny, black structures ( 1 ⁄240 inch; 0.1 mm) 

arising from the stomates on the undersides 

of mottled or brown-tipped needles. 

Figure 1. Black fruiting bodies on the undersides 

of needles pushing up through the 

stomates. Courtesy of Tracey Olson, PDA 

Occasionally, a purple band will be visible 

between the dead tip and green base of the 

needle. These fruiting structures may occur 

on all ages of needles—even on needles 

that appear healthy—so it is important to 

examine many needles on a tree, not just 

symptomatic current-year needles. Fruiting 

bodies found on older needles attached 

to the tree are still capable of releasing 

spores. Infected trees may appear thin and, 

in severe cases, may retain only previous 

year’s growth. Pay particular attention to 

lower and inner branches when scouting for 

this disease. Unfortunately, by the time this 

disease is found, much of the green foliage 

on the tree is infected. Rhabdocline and 

Swiss needle casts can both be found on the 

same tree. 


Like most other fungal diseases, Swiss 

needle cast favors rainy, moist, and cool 

weather. Fruiting bodies develop through 

the stomata on the underside of the needles 

by early winter and are visible from late 

winter into spring, but they only mature 

about the time of bud break (Figs. 1 and 2). 

Fruiting bodies from 1- to 3-year-old needles 

are capable of producing spores, which are 

released from the undersides of needles and 

carried by splashing water or wind to new 

growth, where they penetrate and infect the 

needle (Fig. 3). Though primary infection 

occurs on new growth, occasionally previous 

years’ needles will also become infected at 

this time (Fig. 4). This signifi cantly increases 

the infection potential of this disease. 

Symptoms are much slower to develop and 

may not be apparent for as long as 2 years 

after infection. Spore release continues for 

a longer time than Rhabdocline, the other 

common needle cast disease of Douglas-fi r. 

Figure 2. Fruiting bodies arranged in two rows 

on the undersides of needles. Courtesy of 





Infection 

Symptoms 

Casting 


Bud Break 



Strategies 


• Plant trees with expanded spacing (6 feet 

by 6 feet, or 1.83 by 1.83 m) to allow for 

good air circulation. Avoid planting on a 

north-facing slope, near hardwood forests, 

or in a low, damp area. These types of 

areas generally retain moisture on needles 

for longer period of time. 

• Plant resistant or tolerant tree varieties; 

avoid Douglas-fi rs from Rocky Mountain 

seed sources. 

• Remove and destroy unmanaged Douglasfi 

r that may act as a source of inoculum. 

Preseason 

• Properly water and fertilize to promote 

good health of trees. 

• Maintain proper weed control year-round. 

• Begin scouting for fruiting bodies in 4- to 

10-year-old trees in late winter. Select 20 

or more trees and examine the undersides 

of current and older years’ needles on 

three branches per tree. Select branches 

from the lower third of the tree and look 

at 1- and 2-year-old needles. Pay particular 

attention to trees that appear off color 

or thin. Concentrate scouting on trees 

planted in areas that are more conducive 

for disease development. 

• Scouting can be more effective on an 

overcast day when needle mottling is 

more apparent. 


• Threshold level: Ask state/regional plant 

inspectors about regulatory thresholds. 

• In late April/early May, begin regular 

scouting for bud break in each target 

block. Make the fi rst fungicide application 

when the buds are ½–2 inches (1.27–5.08 

cm). 




Biological 


at this time. 

Mechanical 

• Minimize disease transmission by shearing 

only when trees are dry. 

• Shear healthy trees/blocks fi rst to minimize 

transmission of spores on equipment 

and personnel. 

• Sterilize equipment in contact with the 

disease by soaking in denatured alcohol 

for 3 minutes. 

• Remove and destroy heavily infected trees 

prior to bud break. 

Biorational 


at this time. 

Chemical 

• If treatment is necessary, apply fungicide 

when shoots are ½–2 inches (1.27–5.08 

cm). Make a second application 2–3 

weeks later. A third application may be 

necessary if rainfall is high and temperatures 

remain cool. These treatments 

coincide with the second and third applications 

for Rhabdocline needle cast. 

• Since infected needles may remain 

attached to the tree and continue to 

produce spores for up to 3 years, chemical 

control for 3 consecutive years may be 

necessary. 

• If Rhabdocline needle cast is also present, 

make sure to begin fungicide applications 

at fi rst sign of bud break. 


• Inspect plants/nursery stock; buy from a 

reputable company. 

• Collect seeds from trees showing 

resistance for a seed-collection program. 

Figure 3. Infected needles 

brown from tip down. Courtesy 

of Brian Schildt, PDA 

Figure 4. Previous seasons’ 

infected needles may be cast 

or remain on the tree. Courtesy 

of USDA Forest Service North 

Central Research Station 

Archive, Bugwood.org 

(#1406194) 

SWISS NEEDLE CAST .............................................................................................................................................................................................................................................................................. 71

SHOOT AND BRANCH INJURY 

Diseases, insects, and mite pests that kill 

or cause serious damage to new growth 

and entire branches are discussed on 

pages 73–104. The damage may distort 

or kill the shoot or branch but usually 

does not result in death of the entire 

tree. 

SHOOT AND BRANCH INJURY .............................................................................................................................................................................................................................................................. 72

Hosts 

• Pines: Austrian, Scotch, eastern white, 

Jack, red, lodgepole, Ponderosa, Virginia, 

and others 


• Moderate–severe 


• Small, elliptical cankers at bases of 

needles 

• Resin droplets at canker site 

• Clusters of small, black fruiting bodies 

on cankered tissue 

• Flagged (dead and brown) branches 

throughout tree 

• Sapwood beneath canker stained gray 

to black 


• Frost damage 

• Pales weevil adult feeding 

• Diplodia (Sphaeropsis) tip blight 

• White pine blister rust 


Four native North America species of 

Atropellis cause canker on pines. Atropellis 

piniphila and A. pinicola occur in the western 

and northwestern parts of the country, 

respectively. Atropellis apiculata is only found 

in Virginia and North Carolina. The species 

occurring in Pennsylvania is A. tingens. This 

article focuses on A. tingens. 

This fungal disease of pines causes small, 

elliptical, blue-black cankers (or areas of 

dead tissue) about 0.79 inch (2 cm) long 

underneath the bark of twigs and branches, 

originating at needle bases. Small resin 

droplets form on the bark surface around 

the margins of cankers. Multiple cankers 

may join to girdle a twig or branch. Needles 

on these girdled twigs/branches begin to discolor 

and the twig/branch eventually dies. 

Figure 1. Fruiting structures of Atropellis canker 

on an infected Scotch pine branch. Courtesy of 


These fl agged branches are most noticeable 

in spring and early summer. In addition 

to patches of dead branches, this disease 

can also be recognized by the clusters of 

black, cup-shaped fruiting bodies that are 

0.08–0.16 inch (2–4 mm) long and arise on 

the dead bark of 2- to 3-year-old cankers. 

Cutting into cankered areas reveals darkly 

stained sapwood. This distinguishes Atropellis 

canker from Diplodia, which discolors 

the wood reddish brown. 

The death of branches due to A. tingens 

can disfi gure Christmas trees, making them 

less economically desirable. 


Approximately one year after infection 

begins, the fungus develops barely visible, 

round, black structures (stromata) on the 

surface of the bark above the cankers. These 

structures produce noninfectious spores. 

Black, cup-shaped fruiting bodies (apothecia) 

form on the bark surface (Fig. 1) approximately 

2–3 years after infection. Apothecia 

are 0.08–0.16 inch (2–4 mm) in diameter 

and can be seen easily without magnifi cation. 

Beginning in summer, these fruiting bodies 

swell in moist weather conditions and forcefully 

release infectious spores. Spore release 

continues through the early fall. 

The fruiting bodies continue to sporulate 

annually, and will even do so for a short 

time on a standing dead or fallen tree if it 

remains wet. The spores are dispersed by 

wind and may be carried up to 328 feet 

(100 m) from the host plant. Spores that 

come in contact with healthy bark or needle 

scars germinate under moist conditions. The 

fungus enters the tree through the base of 

the needle. Infection occurs in living tissue 

and grows rapidly within the fi rst year or 

two, staining the wood tissue blue black as 

it grows (Fig. 2). Fungus growth slows after 

the fi rst couple of years; after 10 years, it 

becomes inactive. 

Figure 2. Distinctive blue-black stain is 

characteristic of Atropellis infection. Courtesy 


ATROPELLIS 

CANKER 

Atropellis tingens 

Lohman and Cash 

(also A. apiculata 

Lohman, Cash, and 

R. W. Davidson, 

A. pinicola Zeller 

and Goodd, and 

A. piniphila [Weir] 

Lohman and Cash) 

Flagging and girdled 

branches symptomatic of 

Atropellis canker. 


PDA 

ATROPELLIS CANKER ............................................................................................................................................................................................................................................................................. ..................................................................... 71 73



Infection 

Symptoms 

Casting 


Bud Break 



Strategies 


• Plant trees with enough space between 

them to encourage airfl ow. 

• Remove and destroy any overgrown 

pines surrounding the proposed block to 

decrease chance of infection. 

Preseason 

• Maintain plant vigor and health with 

proper fertilization. 

• Control weeds under and around trees 

throughout the year. 

• Scout trees for fl agged branches and apothecia 

on the bark of infected branches. 

• Immediately prune and destroy any 

infected material found. Remove and 

destroy any tree with multiple cankers. 


• Continue to scout throughout the year 

and prune out and burn infected material. 




Biological 


at this time. 

Mechanical 

• Prune cankered branches 6–12 inches 

(15.2–30.5 cm) below the canker or 

where the branch attaches to the main 

stem. Remove and burn infected material. 

Disinfect shears with 70 percent alcohol 

or a bleach solution between cuts, as 

spores can be spread on tools. 

• Remove and burn trees with cankers 

on the main stem or trees with a heavy 

infection. 

Biorational 


at this time. 

Chemical 


at this time. 


• Buy and plant disease-free stock only. 


Hosts 

• All true fi rs 


• High 

Symptoms 

• Flat top or crooked terminal 

• Gouting (swelling) around buds and 

internodes 

• Stiff, infl exible trunk and large lateral 

branches 

• White, cottony masses on trunk and 

large branches 

• Dead shoots or branches (red or brown 

needles) 


• Phytophthora root rot 


Balsam woolly adelgid females are softbodied, 

spherical, purplish-black, wingless 

insects. They are about 1 ⁄25 inch long (< 1 

mm) and are not mobile. During the winter, 

immature nymphs can be found on bark. 

They are dark and have white, waxy rods 

down their backs and around the edges 

of their bodies. At this stage, they closely 

resemble the eggs of balsam twig aphid. As 

the mature, they continue to secrete this 

waxy substance, which gives them a covering 

that may cause them to resemble minute 

cotton balls by the time females are present. 

Eggs are oblong and deposited in a cluster 

behind the female. The orange-brown 

crawlers are the only stage with functional 

legs. Red eyespots are also visible on the 

crawlers. A hand lens is needed to effectively 

locate all stages. 

Symptoms can be helpful in detecting 

balsam woolly adelgid, but it may take several 

months for damage to become visible. 

Symptoms develop slowly since the insect 

only feeds on the bark, not the needles. 

When a tree begins to show symptoms, 

it means that the tree has been infected 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

since before bud break of that growing 

season. The fi rst most noticeable symptom 

is a fl at top or weak, crooked terminal. As 

the infestation progresses, trees may have 

swelling, or gouting, around the buds and 

shoot nodes. The swollen growth is a result 

of a chemical inserted as the adelgid feeds 

on the branches. When adelgids feed on the 

main trunk, they cause the tree to become 

brittle due to the formation of scar tissue. 

When harvesting, examine stumps for this 

dark ring of wood. 


No male balsam woolly adelgids are present 

in North America. Females reproduce 

parthenogenetically (by producing eggs 

that are exact clones of the adult). The 

immature nymphs overwinter on the trunk, 

larger branches, and around buds. In early 

spring, they mature and the female adelgids 

are present. Females cover themselves 

with waxy, wool-like covering—hence 

the name balsam woolly adelgid (Fig. 1). 

Figure 1. Adelgids covered with a protective 

white, woolly wax. Courtesy of USDA Forest 

Service Region 8 Archive, Bugwood.org 

(#1510053) 


Apply horicultural oil. 

BALSAM 

WOOLLY 

ADELGID 

Adelges piceae 

(Ratzeburg) 

Swelling, or gouting, on 

branches resulting from 

abnormal cell growth 

caused by adelgid chemical 

secretion. Courtesy of USDA 

Forest Service Region 8 


(#1510051) 

BALSAM WOOLLY ADELGID ................................................................................................................................................................................................................................................................. ..................................................................... 71 75

Figure 2. Cluster of eggs produced 

by females under white, 

woolly wax. Courtesy of Scott 

Tunnock, USDA Forest Service, 

Bugwood.org (#2252066b) 

Each female produces a cluster of eggs 

under the woolly mass surrounding her body 

(Fig. 2). Up to 200 eggs can be produced in 

the laboratory, but in the fi eld the number 

of eggs is reduced and depends on the condition 

of the host tree. Eggs hatch in about a 

month, around the time of bud break. 

When crawlers emerge, they can live 

up to 2 days while searching for a place 

to settle. Since crawlers lack mouthparts, 

they must molt before feeding can begin. 

These nymphs insert their piercing-sucking 

mouthparts into the bark to feed, but, once 

inserted, the adelgid is not able to move. 

While feeding, adelgids secrete a chemical 

that causes the abnormal growth of cells in 

the area. This leads to swelling or gouting, 

branch dieback, and possible death of the 

tree (Fig. 3). The stationary crawlers will 

molt several times before becoming adults. 

In midsummer, these adults lay eggs, which 

again hatch into crawlers. The crawlers 

overwinter and start the cycle again the following 

spring (Fig. 4). In Pennsylvania, two 

generations occur per year. 

Figure 3. Gouting, or swelling, from growth 

of cells in the area. Courtesy of Robert L. 

Anderson, USDA Forest Service, Bugwood.org 

(#1748038) 

Figure 4. Overwintering balsam woolly adelgid 

nymphs. Courtesy of Rayanne D. Lehman, PDA 


Strategies 


• Remove and destroy unmanaged fi r that 

may act as a source of inoculum. 

• Purchase and plant pest-free stock from a 

reputable nursery. 

Preseason 

• Scout at waist height for adelgid symptoms 

on branches (swollen areas) and 

main trunk (white, cottony masses). Look 

for fl at tops on trees or dark reddish rings 

in wood of cut stumps. 


• Maintain good weed control. 

• Butt-prune trees when appropriate. 

• Scout for adult adelgids in late spring 

through late summer. Look for the white, 

waxy, wool-like covering, remove suspected 

area, and examine it with a hand lens 

or under a microscope to confi rm adelgid 

infestation. 

• Tag any infested or suspected trees. 

• Scout for terminal damage in fall after tree 

growth has ended for the season. Look 

for fl at tops by walking back and forth 

through rows spaced 6–10 feet (1.83–3.05 

m) apart. If a suspect tree is found, rock 

the tree back and forth to check for a stiff 

trunk. If not confi rmed, fl ag the tree and 

recheck in about a month. 

• If the adelgid is found, treat the fi eld 

before bud break of the following season. 

Treat after harvesting any trees from 

fi eld and possibly time with other pest 

treatments (e.g., with balsam twig aphid 

control). 

• Examine stumps from the current season 

for evidence of scar tissue. 





Biological 

• Several foreign predators have been introduced, 

although the level of control they 

provide is still unclear. Aphidecta obliterate 

(L.), Laricobius erichsonii (Rosenhauer), 

and Pullus impexus (Mulsant) are beetles. 

Aphidoletes thompsoni Mohn, Cremifania 

nigrocellulata (Czerny), and Leucopis obscura 

Haliday are fl ies. 

Mechanical 

• Cut and burn heavily infested trees that 

will not be salable. Do not cut during 

crawler activity. 

• Clear-cut infested blocks. 

Biorational 

• Horticultural oil spray: apply thoroughly 

to the trunk and bark of large branches 

in late fall, winter, or early spring; oil can 

damage foliage if not applied properly. 

— Only apply oil when temperatures are 

above freezing. 

— Oil will remove “bloom,” or blue color, 

from blue specimens. 

Chemical 

• Use a high-pressure sprayer to apply an 

appropriate insecticide during the fi rstgeneration-crawler 

stage; fully cover all 

trees to achieve the best control. Note: 

Control for balsam woolly adelgid may 

kill natural predators of other pests. 

• Scout one month after treatment to make 

sure adelgids are dead; look for new woolly 

spots. 


• Scout every year to determine if and when 

treatment or re-treatment is necessary. 

BALSAM WOOLLY ADELGID .................................................................................................................................................................................................................................................................. 77

BOTRYTIS 

BLIGHT 

Botrytis cinerea 

Pers.: Fr. 

New growth infected with 

Botrytis blight. Courtesy of 


Hosts 

• All Christmas tree species 


• Low 


• Spots on needles and shoots that appear 

water soaked 

• Blighted new growth that resembles frost 

damage 

• Brown lesions on needles and shoots that 

can grow and girdle the shoots 

• Gray, fuzzy, fungus structure on infected 

shoots and needles 

• When the fungus is dry, clouds of spores 

will be released when disturbed 


• Atropellis canker 

• Diplodia (Sphaeropsis) tip blight 

• Cytospora canker 

• Frost damage 

• Pales weevil adult feeding 


This fungal disease can infect hundreds of 

woody and herbaceous plants, including most 

species of trees grown as Christmas trees. 

This fungal disease does not produce typical 

fruiting structures as seen in other conifer diseases. 

Instead, Botrytis cinerea, or gray mold, 

produces a gray, weblike mycelium over the 

surface of the diseased area. Spores are borne 

on stalks on this hairlike growth. 

Symptoms appear in spring and may 

follow frost damage or an extended period of 

shoot elongation due to cool temperatures. 

Infection will not occur on woody tissue, 

but the succulent new growth is susceptible. 

Infected tissue may appear water soaked 

and will eventually turn brown (Fig. 1). 

Figure 1. New shoot browned from Botrytis 

infection. Courtesy of Tracey Olson, PDA 

Figure 2. Dieback of new growth caused by 

Botrytis. Courtesy of Rayanne D. Lehman, PDA 

Tip dieback may occur when the infected 

areas girdle the shoots (Fig. 2). The disease 

develops under humid conditions with cool 

temperatures. Seedling beds are generally 

most susceptible to this blight because of 

the tight spacing. 


Botrytis overwinters in the Northeast in 

moist, decaying plant tissue either as mycelium 

or as small, black, resting structures, 

or sclerotia. As the temperatures warm in 

spring, these resting structures start to germinate 

and threadlike mycelium will begin 

to cover the diseased tissue (Fig. 3). Massive 

amounts of spores are formed on this 

mycelium, especially in the morning hours. 

They are spread to healthy tissue by wind 

or splashing water. When the spores come 

in contact with new plant tissue and during 

periods of high humidity or moisture, they 

will begin to germinate and penetrate the 

plant tissue. Spores can be produced continuously 

during moist conditions and may 

remain dormant for up to 3 weeks before 

they germinate. 

Figure 3. Threadlike mycelium with spores 

covering infected tissue. Courtesy of 


SHOOT AND BRANCH INJURY ........................ 

.............................................................................................................................................................................................................................................................. 78

This fungus can persist in either dead or 

living plant tissue. Infection typically takes 

place in shaded areas with high humidity, 

such as where trees are planted close 

together and on the lower branches of 

trees. A growing season that has an early 

warm spell, causing bud and shoot growth 

to begin, followed by an extended cool/ 

damp period, which suspends needle and 

shoot development, is especially favorable 

for Botrytis. This fungus is a relatively weak 

pathogen. New shoots that are injured by 

frost or mechanical means are most prone to 

this disease. 


Strategies 


• Space trees to encourage drying. 

• Control weeds to reduce the moisture 

level around the trees. 

Preseason 

• Promote tree health by using good cultural 

practices, such as proper fertilization 

and adequate watering. 


• Scout trees that are densely planted and 

shaded. Look for tree shoots that appear 

rain soaked or exhibit the gray webbing. 

• Prevent tree injury from machinery. 

• Control damage from insects and other 

diseases. 




Biological 


at this time. 

Mechanical 

• Prune infected tips out of established 

trees after foliage has died. Disinfect 

shears with 70 percent alcohol or a bleach 

solution between cuts since spores can be 

spread on tools. 

• Remove infected seedlings from seedling 

beds to prevent disease spread. 

Biorational 


at this time. 

Chemical 

• In general, fungicide sprays are unnecessary 

for established Christmas tree plantings. 

• For seedling beds, preventative fungicide 

sprays may be helpful in preventing infection 

during a cool, moist growing season. 


• Do not purchase or accept infected 

nursery stock. 

BOTRYTIS BLIGHT ................................................................................................................................................................................................................................................................................... 79

COOLEY 

SPRUCE GALL 

ADELGID ON 

SPRUCE 

Adelges cooleyi 

(Gillette) 

Terminal galls of Cooley 

spruce gall adelgid. Courtesy 

of Cathy Thomas, PDA 

Hosts 

• Colorado blue spruce 

• Occasionally found on other spruces 

• Alternate host: Douglas-fi r (see Needle 

Discoloration and Injury section for 

Cooley spruce gall adelgid on Douglas-fi r 

on page 29) 


• Low 



• Green, purple, or brown, pineapple- or 

pinecone-shaped galls, 1½–2½ inches 

(4–6 cm) long, on the tip of new growth 


• None; galls may be mistaken for cones 


The Cooley spruce gall adelgid is a small, 

soft-bodied, black insect that may be winged 

or wingless. It possesses piercing-sucking 

mouthparts to suck sap from its host. On 

spruce trees, its presence can be identifi ed 

by the pineapple- or pinecone-shaped swelling 

or gall. This gall is only found at the tip 

of new growth. Galls are green or purple in 

color and turn brown as they dry out in the 

summer. When dried, they may be confused 

with cones. 


nymph in bark crevice. 


PDA 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 


The Cooley spruce gall adelgid overwinters 

as immature females. On spruce, these overwintering 

forms can be found in the bark 

crevices of twigs (Fig. 1), just below the buds 

or around the base of the buds. In March and 

April, the overwintering immatures feed on 

sap and gradually swell. They secrete long, 

waxy fi laments over their bodies as they 

mature (Fig. 2). Shortly before bud break, 

each female produces a cluster of 150–200 

eggs inside this mass (Fig. 3). The brownish 

nymphs emerge from the eggs as the spruce 

buds are opening. They immediately crawl 

into the bud and begin feeding on the 

elongating needles. 

By the time eggs hatch, the bases of 

the newly expanding needles have already 

started to swell due to feeding by the overwintering 

form. When the newly emerged 

nymphs feed, they inject a toxin through 

their saliva that causes further swelling of the 

needle bases. Eventually, the needle bases 

fuse together, forming a terminal gall (Fig. 

4). Each nymph is contained in an individual 

chamber, where they will continue to feed on 

the succulent tissue inside the gall (Fig. 5). 

The gall turns brown and opens in 

midsummer, and the mature nymphs emerge 

(Fig. 6). They move to the needles, molt 

into winged adults, and will either remain 

on the spruce or fl y to a Douglas-fi r tree 


Spray must occur before nymphs wax over. 

Figure 2. Nymphs covered 

with waxy fi laments. Courtesy 

of PDA 

Figure 3. Cluster of eggs laid 

under protective waxy threads. 

Courtesy of Whitney Cranshaw, 

Colorado State University, 



.............................................................................................................................................................................................................................................................. 80

Figure 4. Fused 

needle bases 

forming a 

terminal gall. 

Courtesy of 

Cathy Thomas, 

PDA 

Figure 5. 

Adelgids 

growing in 

chambers 

inside a gall. 

Courtesy of 

Sandy 

Gardosik, PDA 

(Fig. 7). (See Cooley Adelgid on Douglasfi 

r for life history of that host.) In fall, 

winged adults from Douglas-fi r may travel 

back to the spruce tree to produce another, 

nondamaging generation. This fall generation, 

which includes both male and female 

adelgids, will produce the overwintering 

forms by mid-October. While the Cooley 

spruce gall adelgid alternates its life cycle on 

two different hosts, it does not require both 

hosts to complete its life cycle. It may cycle 

continuously on spruce or Douglas-fi r. 


Strategies 


• When planting, separate Colorado blue 

spruce and Douglas-fi r trees; this will not 

eliminate problems, but it may help lessen 

the severity. 

Preseason 

• Scout for brown galls in late fall, winter, 

and early spring to note which trees had 

a problem. 

• Scout for overwintering nymphs using 

a 15X hand lens. 


• Growing degree days: Recommended 

control against nymphs should occur 

in the spring as nymphs begin to swell, 

before waxing over, at 22–81 GDDs and 

then in the fall at 2,800–3,000 GDDs. 

• Threshold level: Treatment is warranted 

when 5 percent of trees have 10 or more 

galls. 




Biological 

• Encourage natural predators such as lacewings, 

predatory bugs, and fungi. Refer to 

Appendix B: Biological Controls Photo 

Chart for pictures. 

Mechanical 

• Pruning: Remove and destroy unopened 

green galls on trees before the release of 

mature nymphs (usually mid-July). 

Biorational 

• Dormant oil: To control overwintering 

nymphs, apply in early spring or late fall 

(late October/early November) when trees 

are not actively growing. 





Chemical 

• Spring insecticide: Apply in mid-April 

when greenish-black adelgids are still 

exposed, before waxy fi laments cover the 

immature females, and eggs are present; 

temperature should be above freezing. 

• Fall insecticide: Apply in late September 

through October; thoroughly cover 

branches and buds. This is the most 

reliable time to spray. 

• A general rule for fall application is to 

wait until after the fi rst frost to guarantee 

that nymphs will be settled when the 

spray is applied. 


• Only plant pest-free trees from a reputable 

source. 

Figure 6. Browned galls after 

release of adelgids. Courtesy of 

Whitney Cranshaw, Colorado 

State University, Bugwood.org 

(#5369749) 

Figure 7. Winged adult adelgid. 




COOLEY SPRUCE GALL ADELGID ON SPRUCE ...................................................................................................................................................................................................................................... 81

DIPLODIA 

(SPHAEROPSIS) 

TIP BLIGHT OF 

PINES AND 

OTHER 

CONIFERS 

Diplodia pinea (Desm.) 

J. Kickx f. (formerly 

Sphaeropsis sapinea) 

Shoots turning yellow green 

and then brown from 

Diplodia. Courtesy of 


Hosts 

• Austrian, red, Scotch, and other two- or 

three-needled pines 

• Rarely on Douglas-fi r and spruce 




• Brown, yellow, gray, or straw-colored 

needles at tip of current season’s growth; 

needles usually stunted and shoot may 

curl 

• Branch dieback 

• Small, black fruiting bodies on needles, 

cones, or shoot tissue 

• Cankers on stems or branches 

• Oozing resin that adheres to blighted 

needles 


• Pales weevil feeding 

• European pine shoot moth 

• Nantucket pine tip moth 

• Pine shoot beetle 

• Scleroderris canker 

• Winter drying 

• Drought 


Diplodia (formerly Sphaeropsis) tip blight 

is caused by the fungus Diplodia pinea. It 

is the most common and severe disease of 

pines in Pennsylvania and attacks trees of 

all ages. Tips of infected current-year shoots 

will blight. Needles on these dead tips are 

usually tan to straw colored, shorter than 

normal, and typically remain attached. 

Figure 1. Mature fruiting bodies embedded in 

the needles. Courtesy of Joseph O’Brien, USDA 


When the fungus attacks larger branches or 

the main trunk through wounds, misshapen 

tops and even branch or tree death can 

result. Cankers are generally found only on 

mature trees, while tip blight attacks trees of 

any age. 

Trees are most susceptible to infection 

from the time buds begin to open until 

the needles are fully elongated. Once the 

fungus enters the needle, it kills that tissue 

very quickly. Symptoms are visible on the 

current season’s growth and on second-year 

cones. Infected needles will be stunted, straw 

colored, and occasionally glued together with 

hardened resin. Cankers may be found on the 

fi rst branch whorl of infected twigs. Beginning 

in late summer, minute, black, fruiting 

bodies (pycnidia) are visible with a 10X 

hand lens on needles, cones, and tissue found 

attached or unattached to the tree. Woody 

tissue beneath cankers is a light brown to 

amber color when bark is stripped away. If 

the wood beneath these cankers is gray to 

black in color, Atropellis may be present. 


The fungus overwinters in infected needles, 

cones, and woody tissue both on and beneath 

the tree (Fig. 1). During wet weather 

from March through September, the fruiting 

bodies mature and release brown, oval 

spores (Fig. 2). The spores are distributed 

by wind, water, animals, and people to the 

new growth, where they germinate on the 

needles. The fungus enters needles through 

the stomata or may enter branches through 

wounds caused by hail, insects, or pruning. 

The infection reaches the base of the needle 

in a matter of hours, leaving a small, brown 

lesion with a resin drop at the point of entry. 

Figure 2. Fruiting bodies embedded in bark surface 

of twigs. Courtesy of John W. Schwandt, 



.............................................................................................................................................................................................................................................................. 82

The fungus continues to grow into the 

twig and results in browning of the attached 

needles and subsequent cankers 

on the twig (Fig. 3). Needle elongation is 

diminished after infection and dying shoots 

turn yellow-green before becoming straw 

colored. A girdling canker is produced when 

the disease reaches twigs, branches, and the 

main trunk. This canker may also exhibit 

resin fl ow. Tissue above the canker dies and 

major portions of the tree may be killed as 

a result. 

In the second year, cones can become 

infected (Fig. 4). While this does not harm 

the tree in any way, infected cones serve as 

a large reservoir of spores and contribute 

to the spread of the disease. This disease is 

present year-round. 

Douglas-fi r and spruces have occasionally 

been observed with Diplodia tip blight 

(Fig. 5). In most cases, this has been a result 

of unusual circumstances that have high 

disease pressure due to adjacent infected 

pines in windrows or nursery blocks. 


Strategies 


• Plant disease-free stock. 

• Avoid planting susceptible species on sites 

where they may be more prone to insect 

injury, disease, or stressed conditions. 

• Do not plant trees near an area that is 

already in infected with Diplodia. 

Preseason 

• Maintain tree vigor throughout the year 

with adequate water and fertilization. 

• Mow weeds and area around trees to allow 

for air circulation. Avoid mower or string 

trimmer damage. 


• Control insects and other pests to reduce 

stress level of trees and potential infection 

sites from wounds created by feeding. 

• Scouting: 

— Late spring/early summer: Randomly 

select 50 trees (of any age) and look 

for stunted, curled, or dead shoots of 

current-year growth. Tag or mark the 

trees. 

— In fall, scout tagged trees for fruiting 

bodies. Look beneath the fascicle 

sheath of needles that are straw colored 

or held in place with resin. 

— If more than 10 percent of scouted 

trees are unfi t for sale due to Diplodia, 

consider treating the entire plantation 

with fungicide next spring. 




Biological 


at this time. 

Mechanical 

• Prune and remove infected material 

(twigs, branches, cones) during dry weather 

when fruiting bodies are not releasing 

spores. Remove and burn or bury pruned 

material. 

Biorational 


at this time. 

Chemical 

• Apply appropriate fungicide in early 

spring when candles begin to elongate. 

Continue applications at 1- to 2-week 

intervals until needles reach full size 

(usually two to four sprays). 

• Note: Fungicide application will not protect 

seed cones from becoming infected. 

No recommendations are available at this 

time to prevent infection of seed cones. 


• Prevent new foliage from becoming 

infected by using measures listed above. 

Figure 3. Canker causing twig 

girdling and leading to tip 

dieback. Courtesy of Tracey 

Olson, PDA 

Figure 4. Infected second-year 

cones serving as a source of 

infectious spores. Courtesy 

of USDA Forest Service North 

Central Research Station 


(#1406026) 

Figure 5. Diplodia infection on 

Douglas-fi r. Courtesy of Tracey 

Olson, PDA 

DIPLODIA (SPHAEROPSIS) TIP BLIGHT OF PINES AND OTHER CONIFERS ........................................................................................................................................................................................... 83

EASTERN 

PINE WEEVIL 

(FORMERLY 

KNOWN AS 

NORTHERN 

PINE WEEVIL) 

Pissodes nemorensis 

Germar 

Eastern pine weevil adult 

feeding damage. Courtesy 

of Wood Johnson, USDA 

Forest Service, Bugwood.org 

(#1113049) 

Hosts 

• Weakened, dead, or dying trees 

• Pine 

• Occasionally on spruce and Douglas-fi r 




• Discoloration and browning (fl agging) of 

small branches or twigs 

• Small, circular holes in the main trunk or 

base of lower branches 

• Sap excreting from circular holes 

• “Chip cocoons” containing mature larvae 

or pupae under the bark on lower half of 

the tree; not found below root collar area 


• Pales weevil 

• White pine weevil 


Eastern pine weevil adults have long snouts 

typical of all weevils. The body is ½– 5 ⁄8 inch 

(12–16 mm) long and 1 ⁄8–¼ inch (3–6 mm) 

wide. Females are larger than the males. 

The main body is covered with scalelike 

hairs varying from medium brown to almost 

black. The wing coverings have noticeable 

patches of white and gold scales on the 

rear half. Frequently, these patches are not 

joined. 

The legless larvae are white with medium 

brown heads. They have sparse yellow hairs 

and are slightly “C” shaped. A mature larva 

is about ½ inch (12 mm) long. Mature 

larvae and pupae are found in characteristic 

“chip cocoons” under the bark of the tree. 

These structures are composed of excelsiorlike 

strands of wood removed by the mature 

larvae. 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

Eastern pine weevil and white pine 

weevil are largely inseparable based only on 

appearance of the adults, larvae, and pupae. 

The two species are often found together in 

alcohol-and-turpentine-baited pyramidal, or 

Tedder’s, traps used by growers to pinpoint 

the emergence time of white pine weevil. 

Eastern pine weevil is slightly larger in all 

stages, but separation of the two species is 

diffi cult even for an entomologist. However, 

the two species reproduce in different parts 

of the tree, which is a reliable tool for identifi 

cation. White pine weevil will only be 

found above the second whorl of branches. 

Eastern pine weevil is found on the lower 

trunk down to the root collar area as well as 

under the bark of stumps. 


Most eastern pine weevils overwinter as 

adults in the soil at the base of trees but 

are occasionally found near ground level 

in cracks and crevices in the outer bark. 

Very few will overwinter as larvae or pupae 

in the inner bark of infested trees. Adults 

resume activity in April and begin to feed 

on stumps, trunks, and large branches of 

healthy host trees (Fig. 1). They chew a 

small, circular hole in the outer bark and 

insert their mouthparts to feed on the inner 

bark. Young trees and seedlings are also 

attractive hosts in the spring and may be 

killed by this feeding. 

Figure 1. 

Adult 

feeding on 

a stump. 

Courtesy of 

Rayanne D. 

Lehman, PDA 



.............................................................................................................................................................................................................................................................. 84

After continuously feeding for about 

3 weeks, the weevils move to appropriate 

breeding sites to mate and lay eggs. These 

sites include fallen trees, dying or dead trees, 

stumps, and sometimes trunks of stressed 

transplants. Females deposit eggs singly in 

holes chewed in the bark, preferring branch 

nodes to open bark areas. Eggs are commonly 

deposited on the lower portions of the tree, 

often in the shadiest spot. Oviposition is 

complete by midsummer, around the time 

the adults die. Eggs hatch in about 8 days 

and young larvae immediately begin feeding 

in the cambial layer (Fig. 2). They will 

continue to feed along the grain of the wood, 

making galleries in the inner bark. Prior to 

pupation, larvae remove wood fi bers from 

the xylem to create a chip cocoon in the 

outer bark (Fig. 3). If they pupate in small 

transplants, the chip cocoons will be in the 

inner wood (Fig. 4). Larvae and pupae are 

never found in the leader (unlike the white 

pine weevil) and never below the soil line 

(unlike the pine root collar weevil). Larvae 

are commonly found throughout the growing 

season. 

The majority of adults emerge 2 months 

after eggs are laid. Adult emergence starts 

in early July and can span several months 

due to the long oviposition period and 

occasional immature stages that overwinter. 

Most eastern pine weevils complete a 

generation in 1 year, but a few individuals 

will require 2 years. Following emergence, 

adults feed briefl y on the bark of available 

stumps, healthy trees, and seedlings before 

overwintering (Fig. 5). 

Figure 2. Larva feeding under the bark of a 

dying tree. Courtesy of PDA 


Strategies 


• Delay planting for 2 years in an area 

where pines were recently harvested. 

• Remove and destroy weakened pines in 

perimeter of block. 

Preseason 

• Remove and destroy stumps. 

~or~ 

• Treat stumps from the previous season’s 

harvest before adult weevils become 

active in spring. 

• Growing degree days: Overwintering 

adults become active from 7 to 100 GDDs. 

• Destroy cull piles and weakened or dying 

trees before weevils become active in 

spring. These weevils are attracted to the 

resins in stumps and slash and will fl y up 

to several miles to seek breeding material. 


• Threshold level: Treat a plantation with 

fi ve or more fl agged tips per tree. 




Biological 


at this time. 

Mechanical 

• Remove dying or dead trees in early spring 

since they are good oviposition sites. 

Biorational 


at this time. 

Chemical 

• In early spring, treat freshly cut stumps 

and the surrounding soil with a registered 

insecticide to control ovipositing adults. 

• If damage is severe, apply foliar sprays in 

late summer to control adults feeding on 

mature trees, transplants, and seedlings. 


• Remove any damaged or highly susceptible 

trees surrounding the plantation 

before planting. 

Figure 3. Larvae in a chip 

cocoon. Courtesy of Gerald J. 

Lenhard, Louisiana State 

University, Bugwood.org 

(#0014079) 

Figure 4. Pupa in a chip cocoon. 

Courtesy of Gerald J. Lenhard, 

Louisiana State University, 


Figure 5. Adult feeding on a 

twig. Courtesy of Rayanne D. 

Lehman, PDA 

EASTERN PINE WEEVIL ........................................................................................................................................................................................................................................................................... 85

EASTERN 

SPRUCE GALL 

ADELGID 

Adelges abietis 

(Linnaeus) 

Adelgid nymphs maturing 

inside a protective gall. 


PDA 

Hosts 

• Norway spruce (Picea abies) 

• White spruce (P. glauca) 


• Moderate 


Before and at Bud Break 

• Stem mothers and small masses of eggs 

covered with white, woolly wax 


• Green- or brown-colored galls at the base 

of new shoots; growth beyond gall may be 

dead or stunted 


• Cooley spruce gall adelgid (galls will be 

found at tips of new growth) 


This insect is diffi cult to fi nd but can easily 

be identifi ed by the gall it forms. In spring 

as the new growth elongates, galls resembling 

small pineapples, ½ –1 inch (1.0–2.5 

cm) long, are formed at the base of the 

new growth—in contrast to Cooley spruce 

galls, which form at the end of the new 

shoot. New growth will continue beyond 

the gall, somewhat obscuring the gall from 

view. Initially, galls are green, but they turn 

brown and open in late summer, releasing 

the nymphs. Both previous and current-year 

galls may be visible on trees. The galls may 

cause stunting and death of shoots, resulting 

in disfi gurement of the branches and tree, 

but they rarely kill the tree. 

Overwintering immature females can be 

found from fall through early spring at the 

base of the terminal and lateral buds and in 

bark crevices of twigs. They are dark and 

may have a faint ring of white, waxy material 

surrounding their bodies. As bud break 

approaches, they will increase the waxy 

material before depositing eggs. A 15X hand 

lens is required to locate overwintering 

eastern spruce gall adelgids. 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 


The eastern spruce gall adelgid overwinters 

as an immature female called a stem mother 

(Fig. 1). As bud break approaches, the stem 

mother matures and secretes white, woolly 

wax around her body (Fig. 2). She deposits 

100–200 greenish-brown eggs in this mass 

(Fig. 3). 

Figure 1. Overwintering immature 

females (stem mothers). Courtesy of 


Figure 2. Female secreting waxy 

fi laments to cover her body. Courtesy 

of PDA 

Figure 3. Eggs under white, woolly 

wax made by the stem mother. 




.............................................................................................................................................................................................................................................................. 86

At bud break the eggs hatch into 

yellowish-colored nymphs, which move to 

the new growth and begin feeding. This 

feeding causes abnormal twig growth, which 

results in gall formation (Figs. 4 and 5). 

Figure 4. Purplish growth indicating the start 

of a gall on new growth. Courtesy of Tracey 

Olson, PDA 

Figure 5. Gall of eastern spruce gall adelgid. 

Courtesy of Minnesota Department of 

Natural Resources Archive, Bugwood.org 

(#4212052) 

The adelgids remain safe in the gall 

until August/September when the gall turns 

brown and opens to release mature nymphs 

(Figs. 6 and 7). These nymphs mature to 

winged, egg-laying females; there are no 

males. These females generally remain on 

the same tree, depositing their eggs near the 

tips of needles. These second-generation 

eggs result in immature females, which will 

overwinter near buds (Fig. 8). 


Strategies 


• Plant tree varieties that are resistant to or 

are not hosts of eastern spruce gall adelgid. 


within and surrounding the plantation. 

Preseason 

• Scout for brown galls on trees from the 

previous year’s infestation; monitor these 

trees closely for exposed stem mothers 

(before covered with waxy fi laments). 

• Before buds double in size and bud scales 

loosen, apply a dormant oil or spring 

insecticide to manage overwintering 

females. 





• Around bud break time, scout for white, 

woolly egg masses near the buds. 


• Threshold level: At this time, no determined 

threshold number exists for the 

eastern spruce gall adelgid. 

• Growing degree days: Recommended 

control against nymphs should occur in 

spring, before nymphs wax over, at 22–170 

GDDs and in fall at 2,800–3,000 GDDs 

• Scout for green galls; remove and destroy 

them whenever possible before they turn 

brown and open in late summer. 




Biological 


at this time. 

Mechanical 

• Remove green galls before they open in 

late summer. 

Biorational 

• Apply dormant oil in spring to manage 

overwintering females (only when 

temperatures are above freezing). 

Chemical 

• Apply a fall insecticide or dormant oil 

from mid-September to October to 

manage overwintering stages. 



source. 

Figure 6. Browned gall after 

release of adelgids. Courtesy 

of E. Bradford Walker, Vermont 

Department of Forests, Parks, 

and Recreation, Bugwood.org 

(#0907022) 

Figure 7. Gall damage on 

Norway spruce. Courtesy of 

Pennsylvania DCNR Forestry 


(#5018059) 


females around buds. Courtesy 


EASTERN SPRUCE GALL ADELGID ......................................................................................................................................................................................................................................................... 87

GALL RUSTS 

Eastern 

(Pine-Oak Gall Rust), 

Cronartium 

quercuum (Berk.) 

Miyabe ex Shirai 

Western 

(Pine-Pine Gall Rust), 

Endocronartium 

harknessii 

(J. P. Moore) 

Y. Hiratsuka 

Galls may produce spores 

each spring for up to 

10 years. Courtesy of 


Hosts 

• Two- and three-needle pines: Scotch, 

Austrian, Jack, Mugo, red, and Virginia 

• Alternate hosts: oaks for eastern gall rust 




• Stunting, deformation, and dieback of 

twigs and branches 

• Woody, globelike galls on branches or 

stems 

• Yellow-orange aecia (fruiting bodies) 

on surface of mature galls in spring 


• None 


Western (pine-pine) gall rust is the more 

common of the two in Pennsylvania, but 

eastern (pine-oak) gall rust can also occur. 

Both produce visible, globe-shaped galls 

on the stems or branches. The galls will 

increase in size each year and can end up 

being several inches in diameter. Around 

mid-May on 2-year-old and older branches, 

the blisterlike membrane of the fruiting 

structures (aecia) ruptures, covering the 

surface with a mass of powdery, yelloworange 

spores (aeciospores). These spores 

are dispersed in the wind and will infect 

new plants. Western gall rust can easily 

re-infect the same tree and may result in 

multiple galls on the same tree and even 

the same branch. If a tree has a signifi cant 

amount of galls, stunting and dieback of 

branches may occur and lead to death of 

the tree. Western gall rust may be especially 

lethal to seedlings. 


Eastern (pine-oak) gall rust requires two 

hosts: pine and oak. The spores released in 

the spring from the galls on pines can only 

infect the leaves of oak, the alternate host. 

Trees in the red and black oak group are 

especially susceptible. Once young leaves 

of oaks have become infected, a repeating 

spore stage occurs and the disease intensifi 

es. By midsummer, spores that are capable 

of infecting only pines are then produced on 

the oak leaves. Absence of oaks in the growing 

area will break the required two-host 

life cycle and minimize the threat of this 

disease. Western (pine-pine) gall rust does 

not require an alternate host and is spread 

directly from one pine to another, infecting 

only new growth during the spring sporulation 

period. 

For both diseases, spores are wind dispersed 

and infect new needles. The fungus 

grows into the shoots, where it establishes 

in the woody tissue of branches (Fig. 1) and 

possibly the main trunk (Fig. 2). Small galls 

may be visible after the fi rst year; however, 

it may take up to 3 years for them to become 

large enough to be noticed (Fig. 3). 

Figure 1. Western gall rust on a branch (not 

sporulating). Courtesy of Susan K. Hagle, USDA 


Figure 2. Gall rust on the main trunk. Courtesy 

of Joseph O’Brien, USDA Forest Service, 


Figure 3. Gall forming at an infection site. 



.............................................................................................................................................................................................................................................................. 88

Galls are formed due to an increase in 

wood production at the infection site. After 

the second year of infection, yellow-orange 

fruiting structures (aecia) will form on 

the surface of the gall (Fig. 4). Galls may 

actively produce spores each spring for up 

to 10 years. 

Figure 4. Powdery, yellow-orange spores on the 

surface of a gall. Courtesy of Tracey Olson, PDA 


Strategies 


• Avoid planting in fi elds adjacent to forest 

stands of oak or pines that show evidence 

of infection. 

• Plant disease-free nursery stock from a 

reputable source. 

Preseason 

• Inspect seedling beds one year after 

planting. Remove any seedlings showing 

signs of galls. 

• In older plantings, remove branches 

with galls prior to active sporulation in 

mid-May. 

• If a tree is heavily infected with multiple 

galls or trunk dieback is visible, completely 

remove and destroy the tree before 

sporulation. 


• Scout fi elds to look for trees with branch 

and stem galls. For those that cannot be 

removed, monitor for presences of fruiting 

bodies. 




Biological 


at this time. 

Mechanical 

• Remove seedlings exhibiting galls from 

the fi eld or cut out branches with galls 

from older, established trees. Do not move 

during period of spore production. 

Biorational 


at this time. 

Chemical 

• For western (pine-pine) gall rust, apply 

a registered fungicide to new growth of 

pines at the time of bud break to prevent 

infection. This must be applied before the 

fungus sporulates and releases the orange 

spores. 


• Buy disease-free and/or resistant stock 

from a reputable nursery. 

• Rotate crop in the planting block to a 

nonsusceptible tree variety. 

GALL RUSTS ............................................................................................................................................................................................................................................................................................ 89

PALES WEEVIL 

Hylobius pales (Herbst) 

Flagged branches are a 

symptom of pales weevil 

feeding. Courtesy of Sandy 

Gardosik, PDA 

Hosts 

• Pines preferred, Scotch more so than 

eastern white 

• Occasionally on Douglas-fi r, fi r, spruce, 

and other conifer species 




• Dead seedlings with missing bark near 

soil line 

• Small holes or pits in the bark from 

feeding that may fi ll with resin if 

infestation is light 

• Yellow needles at branch tips that turn 

reddish brown, giving the tree a “fl agged” 

appearance; bark will be missing behind 

the dead tissue 

• White, C-shaped larvae under the bark of 

stumps and roots 

• Adult weevils in the leaf litter surrounding 

trees or on stumps in early spring 


• Eastern pine shoot borer 

• Diplodia (Sphaeropsis) shoot blight 

• Northern pine weevil 


• Pine shoot beetle 

• Wood borers 

• Bark beetles 


The adult pales weevil is a large, robust 

insect measuring ¼– 2 ⁄5 inch (6–10 mm) 

long with a prominent snout. The weevil is 

black to dark reddish brown in color with 

very small patches of yellow-white scales 

on its head and wing coverings. In early 

spring, adults are frequently found on and 

around 1- to 2-year-old stumps. The larvae 

are cream colored, legless, and “C” shaped. 

They have a light brown head capsule and 

are found burrowed into the cambium tissue 

of stumps and roots. 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

Adult pales weevils are particularly 

damaging to eastern white pine, Douglas-fi r, 

and true fi rs. The weevils feed on terminal 

shoots, consuming the bark down to the 

wood and girdling and killing the shoot. 

This fl agging can seriously disfi gure marketable 

trees (Fig. 1). Larval feeding is generally 

restricted to 1- to 2-year-old stumps and dying 

trees of Scotch, Austrian, and red pine. 

Larvae and pupae may be found on larger 

roots below the soil surface. Larvae pupate in 

chambers plugged with wood fi bers. 

Figure 1. 

Evidence of 

weevil feeding 

indicated 

by fl agging. 

Courtesy of 

Rayanne D. 

Lehman, PDA 


Pales weevils overwinter as adults in the soil 

and become active between April and June 

(Fig. 2). They feed for several weeks on seedlings 

and the tender bark on twigs of more 

mature trees (Figs. 3 and 4). Feeding occurs 

at night, while the weevils take refuge in the 

leaf litter and under logs during the day. 

Figure 2. Adult pales weevil feeding on 

bark of a twig after spring emergence. 




.............................................................................................................................................................................................................................................................. 90

After feeding, the weevils mate and 

females lay pearly white eggs in the subterranean 

roots of pine stumps or dying trees. 

Eggs hatch within 10–14 days and the larvae 

feed beneath the bark until they mature 

in early fall (Fig. 5).The larvae pupate in 

a chip cocoon (Fig. 6) before emerging as 

adults in August–October. These adults feed 

for 2–3 weeks before seeking overwintering 

sites in the soil around fresh-cut stumps or 

beneath dying trees. One generation occurs 

per year. 


Strategies 


• Delay planting for 2 years in an area 

where pines were recently harvested. 

Preseason 

• Remove and destroy stumps. 

~or~ 

Figure 3. Weevil 

feeding damage 

on eastern 

white pine 

branches. 

Courtesy of 

Brian Schildt, 

PDA 

Figure 4. Weevil 

feeding damage 

on Fraser fi r 

leader. 

Courtesy of 

Rayanne D. 

Lehman, PDA 

• Treat stumps from the previous season’s 

harvest before adult weevils become 

active in spring. 

• Destroy culled tree piles before pales 

weevils become active in spring. These 

weevils are attracted to the resins and will 

fl y up to several miles to seek breeding 

material. Traps deployed to monitor for 

white pine weevil will also attract pales 

weevil adults. 

• Growing degree days: Treat stumps 

with insecticide to prevent egg laying 

at 7–121 GDDs. 


• Scout for the presence of different life 

stages of the weevil. 

— For adults: Lay a sheet out under a tree 

after dark and shake the tree. Look for 

weevils that fall from the tree to the 

sheet. 

— For larvae and pupae: Use a knife to 

cut back the bark on stumps or the 

base of dead trees to look for larvae 

burrowing through the tissue or pupae 

in chip cocoons. 

• Scout 50 random trees of any age for 

seedling injury and branch fl agging. 

— If seedlings show any weevil injury 

or older trees average more than fi ve 

fl agged branches, consider treating the 

entire plantation. 




Biological 


at this time. 

Mechanical 

• Remove and burn or chip stumps and 

debris. 

Biorational 


at this time. 

Chemical 

• Dip new seedlings (aboveground portion) 

in appropriate insecticide before planting. 

• In early spring, apply an approved pesticide 

to stumps from previous cutting 

season. Stumps older than 2 years do not 

need to be treated. 

• If a population is extremely high, apply 

an appropriate insecticide to mature trees 

in spring between April and May and 

again in August and September to kill 

bark-feeding weevils. 


• Remove cut stumps or treat stumps with 

an insecticide before replanting a fi eld. 

Figure 5. Larvae feeding under 

the bark of a stump. Courtesy 


Figure 6. Chip cocoon created 

by larvae before pupating into 

adults. Courtesy of PDA 

PALES WEEVIL ......................................................................................................................................................................................................................................................................................... 91

PINE BARK 

ADELGID 

Pineus strobi (Hartig) 

Tree with pine bark adelgid 

infestation on the branches 

and trunk. Courtesy of Brian 

Schildt, PDA 

Hosts 

• Eastern white pine 

• Sometimes on Austrian and Scotch pine 


• Low–moderate (pest seems to have decreased 

since appearance of multicolored 

Asian lady beetle) 


• Small clumps of white, woolly wax on 

candles, branches, or trunks of discolored, 

stunted, weakened, or dying trees 

• Yellow- or purple-colored insects under 

woolly wax 

• Dark blue to green, wool-covered nymphs 

on growing branches (in May–June) 


• Balsam woolly adelgid 


A tree heavily infested with pine bark 

adelgid may have the look of snow on its 

branches and trunk (Fig. 1). However, in 

Christmas tree production, infestations 

seldom reach this stage. Instead, buds, elongating 

candles, and needle bases will have 

the white, woolly mass if pine bark adelgid 

is present. This mass may contain a nymph, 

a wingless, purplish adult female (about 1 ⁄32 

inch long), or up to 25 light yellow-brown 

eggs. Winged forms may be present during 

the growing season. 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

Figure 1. Tree 

with severe 

adelgid 

infestation. 

Courtesy of 

USDA 

Forest Service 

Northeastern 

Area Archive, 

Bugwood.org 

(#1396093) 


Pine bark adelgids overwinter predominately 

as immature females (Fig. 2). By early 

spring when temperatures reach 50°F, the 

female is mature and begins to produce a 

coating of woolly wax (Fig. 3). She deposits 

about 25 eggs under the woolly mass before 

dying. When the crawlers emerge from 

the eggs, they move to a new location and 

begin to feed. Once they have inserted their 

mouthparts into the bark, they are no longer 

capable of movement. 

Both winged and wingless adults result 

from the next several generations. Winged 

forms will fl y to adjacent trees, while wingless 

forms remain on the same trees. Five 

generations are thought to occur each year, 

but all stages may be present at any given 

time during the growing season. 

Figure 2. 

Nymph on elongating 

candle. 

Courtesy of 

Rayanne D. 

Lehman, PDA 

Figure 3. Protective waxy, woolly covering 

produced by mature females. Courtesy of 

Lacy L. Hyche, Auburn University, Bugwood.org 

(#1540233) 



.............................................................................................................................................................................................................................................................. 92


Strategies 


• Do not plant eastern white pine, particularly 

if mature Austrian and Scotch pine 

are growing nearby. 

Preseason 

• Scout for overwintering blue-green 

nymphs under white, woolly wax. 


• Scout for this pest during the entire 

season on susceptible trees of all ages. 

• Growing degree days: Dormant control of 

overwintering immatures should occur at 

22–58 GDDs. 


established. Treat infested trees if adelgids 

are found on numerous shoots or coating 

the bark and predators are not present. 

Many benefi cial insects are attracted to 

pine bark adelgid, so use discretion when 

applying insecticides for control of this 

pest. If possible, spot treat infested trees 

using a backpack sprayer. 




Biological 

• Encourage benefi cial predators such as 

dusty wings, hoverfl ies, and lady beetles 

(Fig. 4). This pest appears to have decreased 

in occurrence and severity since 

the introduction of multicolored Asian 

lady beetle (Harmonia axyridis). 

Mechanical 


at this time. 

Biorational 

• If benefi cial predators are present, use 

insecticidal soap or horticultural oil. 

• Dormant oils: Applications made during 

the fall and early spring that completely 

cover the trunk and branches will kill 

overwintering nymphs. 





Chemical 

• Insecticide: Apply a registered product 

from mid-April to May when adelgids are 

active. 



source. 

Figure 4. Seven-spotted lady 

beetle feeding on pine bark 

adelgid. Courtesy of Sandy 

Gardosik, PDA 

PINE BARK ADELGID .............................................................................................................................................................................................................................................................................. 93

PINE SHOOT 

BEETLE 

(OR COMMON 

PINE SHOOT 

BEETLE) 

Tomicus piniperda 

(Linnaeus) 

Flagging damage from pine 

shoot beetle. Courtesy of 


This is a federally regulated pest. A Compliance 

Agreement is available to enable 

growers within the quarantine area to ship 

trees to other states. Consult your state or 

federal agency for information on the pine 

shoot beetle quarantine area and shipping 

regulations. 

Hosts 

• Pines, especially Austrian and Scotch 

• Occasionally on eastern white pine 




Spring 

• Boring dust (reddish in color) on cut trees 

or stumps 

• Egg and larval galleries beneath the bark 

of cut trees or stumps 

June Through December 

• Shoots on live trees turn yellow or red 

• Drooping shoots and round holes 

surrounded by pitch 

• Shoot completely hollow 


• Other bark beetles 

• Diplodia (Sphaeropsis) shoot blight 


• European pine shoot moth 


Pine shoot beetle eggs are approximately 1 ⁄ 25 

inch (1 mm) long, oval, smooth, and shiny 

white. Larvae are legless, white grubs, up 

to 1 ⁄ 5 inch (5 mm) long, with brown heads. 

Both eggs and larvae are found inside galleries 

under the bark of weakened trees or 

stumps. Reddish sawdust may be apparent as 

the larvae bore in the inner bark. 

Adult beetles are cylindrical, dark brown 

or black, and generally shiny. They can 

be up to 1 ⁄ 5 inch (5 mm) in length. Newly 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

emerged adults are easiest to fi nd in summer 

as they go through their maturation feeding 

stage. These new adults will feed inside 

current-year shoots until fall when they seek 

overwintering sites. They generally feed 

within 6 inches (15 cm) or less of the shoot 

tip, completely hollowing out the shoot. 

This gallery is completely free of sawdust 

and frass, in contrast to the gallery of moth 

larvae. The entrance/exit hole used by the 

adults is round and frequently ringed with 

sap and sawdust. Shoots bored by pine shoot 

beetles turn yellow and may droop or break 

from the tree (Figs. 1 and 2). 

Figure 1. Exit hole from summer 

maturation feeding site. Courtesy of 


Figure 2. Symptom of pine shoot 

beetle infestation. Courtesy of 




.............................................................................................................................................................................................................................................................. 94


Although they can survive as mature larvae 

or pupae, pine shoot beetles most often 

overwinter as adults in the bark or lower 

stems at the base of trees (Fig. 3). In early 

spring, adults become active and fl y to 

recently cut trees and stumps, attracted to 

chemicals released by the trees. Both sexes 

create a club-shaped chamber called a gallery 

between the bark and sapwood (Fig. 

4). The female beetle carves out the gallery, 

which runs parallel with the grain of the 

wood, while the male ejects the debris from 

the tunnel and blocks the opening so other 

males cannot enter. Eggs are deposited in 

niches created along the wall of the gallery 

and hatch 2–3 weeks later. Larvae feed for 

6–8 weeks as they bore horizontally into 

cambial tissues before pupating. 

Figure 3. 

Adult pine 

shoot beetle. 

Courtesy of 

Jim Stimmel, 

PDA, 

Bugwood.org 

(#2117021) 

Figure 4. Adult beetles in a breeding 

gallery. Courtesy of Louis-Michel Nageleisen, 

Département de la Santé des Forêts, Bugwood 

.org (#2101014) 

In early summer, new adults emerge 

from the breeding site and begin feeding 

on current or one-year-old growth of living 

trees. This “maturation feeding” can occur 

from May through October (Figs. 5 and 6). 

Beetles may feed on as many as six shoots 

before overwintering. One generation 

occurs per year. 


Strategies 


• Plant tree species other than pine. 

Preseason 

• Chip or burn any culled pines to remove 

breeding sites. 

• Cut pine stumps low to the ground and 

apply appropriate insecticide to prevent 

larval development in late April/early 

May. 


• Scout for dead or bent shoots on the 

upper half of the tree; shoots may be 

yellow or red and will have small, round 

holes where beetles entered and exited. 

• Clip off and open suspected shoots to 

examine for pine shoot beetle. 



Figure 5. Adult beetle exiting 

from a maturation feeding 

site. Courtesy of Rayanne D. 

Lehman, PDA 

Figure 6. Adult beetle inside 

a feeding gallery within a 

shoot. Courtesy of Rayanne D. 

Lehman, PDA 

PINE SHOOT BEETLE ............................................................................................................................................................................................................................................................................... 95


Biological 


at this time. 

Mechanical 

• Sanitation is critical to controlling 

this pest. 

• Cut stumps less than 4 inches (10 cm) 

from the ground when harvesting trees or 

in the spring before the new generation 

completes development. 

• Chip or burn any unused or discarded 

trees or branches to minimize breeding 

site material. 

• Use trap logs to attract breeding parent 

beetles by systematically placing freshly 

cut pine trees or logs along the edges of 

the fi eld in early spring. The trap logs 

must be chipped or burned after breeding 

occurs but before new adults emerge. 

Specifi c instructions and timetable are 

provided with the federal compliance program 

guidelines. See the USDA-APHIS/ 

PPQ Web site on pine shoot beetle (www 

.aphis.usda.gov/plant_health/plant_pest_ 

info/psb/index.shtml) for further details 

on regulation of this pest. 

Biorational 


at this time. 

Chemical 

• Apply appropriate insecticide to stumps 

to prevent larval development (late April/ 

early May). 

• Apply appropriate insecticide to foliage 

timed with the emergence of the new 

generation of beetles to help manage 

the beetle population at 450–550 GDDs 

(usually early–mid-June). 



source. 


Hosts 

• Two- and three-needled pines, especially 

Scotch 


• Moderate 


• Black sooty mold on surface of bark and 

needles; tree may appear darker from 

distance 

• Clusters of brown and white, helmetshaped 

scales on branches 

• Presence of stinging insects in and 

on trees 

• Reduced plant vigor 


• Pine tortoise scale 

• Cinara aphids 


Note: Many previous reports of pine tortoise 

scale (Toumeyella parvicornis Cockerell) in 

Pennsylvania actually refer to this closely 

related species. Striped pine scale is the more 

common of the two species in Pennsylvania. 

The striped pine scale is a soft scale 

found on the new growth of pines. The 

coverings of adult female scales resemble 

miniature helmets. They are reddish brown 

and may appear to have one or more cream 

or white stripes centrally. This coloration 

varies with age and condition of the scales. 

The coverings of mature females are ¼ inch 

(6.3 mm) long. They are generally found 

in clusters on the outer twigs of host trees. 

Nymphs or crawlers of this scale are tan 

to orange and will be found on the twigs 

and needles of new growth. As this insect 

feeds on the bark of pines, it excretes a 

sugary liquid known as honeydew (Fig. 1). 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

Black sooty mold will grow on the bark and 

needles coated with the honeydew, giving 

the tree a darkened appearance (Fig. 2). 

Frequently, ants, bees, and wasps will be 

found feeding on this honeydew. A heavy 

buildup of sooty mold results in stunting and 

chlorotic growth. 

Figure 1. Striped pine scale excreting a 

drop of honeydew. Courtesy of Brian 

Schildt, PDA 

Figure 2. Black sooty mold development 

resulting from honeydew excretion. 

Courtesy of R. Scott Cameron, Advanced 

Forest Protection, Inc., Bugwood.org 

(#3226096) 


STRIPED 

PINE SCALE 

Toumeyella pini (King) 

Severe scale crawler 

infestation on Scotch pine, 

along with a heavy amount 

of sooty mold (black coating 

on stem and needles). 

Courtesy of Brian Schildt, 

PDA 

STRIPED PINE SCALE .............................................................................................................................................................................................................................................................................. ..................................................................... 71 97

Figure 3. Female striped pine 

scales. Courtesy of Brian 

Schildt, PDA 

Figure 4. Scale crawlers moving 

over the top of a female scale. 


PDA 


Striped pine scale has a single generation 

each year in Pennsylvania. Fertilized, immature 

females can be found on twigs of 

the most recent season’s growth (Fig. 3). 

Occasionally, a few mature females may also 

be seen overwintering for a second season 

on last year’s growth. In early spring, the 

overwintering nymphs resume feeding and 

mature. Females deposit eggs under their 

helmet- or tortoise-shaped covering in early 

summer. These eggs, which may number 

in the hundreds, will hatch over a period 

of several weeks in late June or early July. 

The newly hatched nymphs, or crawlers, are 

very active (Fig. 4). They quickly move to 

the new expanding growth and can even be 

found on the ground under trees. Crawlers 

can attach to insects, birds, animals, people, 

and machinery and be easily moved to uninfested 

trees. Within a day or two, the newly 

emerged crawlers molt, losing their legs and 

gaining long mouthparts. Once they settle 

to feed, this pest is no longer capable of 

moving about on the host. Female scales remain 

on the bark of twigs, while male scales 

settle on the needles. The males are much 

smaller and the winged adults are generally 

unnoticed. 


Strategies 


• Plant trees with adequate spacing between 

them to prevent crawlers from moving 

tree to tree on overlapping branches. 

• Inspect seedling stock before planting for 

presence of scales on the young growth. 

Preseason 

• Scout fi elds of susceptible trees for presence 

of scales on last season’s growth. Pay 

particular attention to trees attractive to 

ants and stinging insects. 

• If only a few trees show severe infestation, 

remove these trees from the fi eld before 

bud break and burn or chip them. 


• Monitor known scale infestation for 

presence of emerging scale nymphs. 

• Growing degree days: Crawlers emerge at 

400–500 GDDs. 

• Avoid mowing during crawler emergence 

to prevent mechanical spread. 




Biological 

• Encourage naturally occurring parasitoids 

and predators. Lacewing larvae and lady 

beetles feed on crawlers. 

• Do not use a broad-spectrum insecticide 

that will kill benefi cial insects. 

Mechanical 

• Remove and burn or destroy severely 

infested trees before bud break. 

Biorational 


at this time. 

Chemical 

• Apply dormant oil in fall or in spring 

before bud break. 





• A single insecticide treatment applied at 

crawler emergence in early summer is generally 

effective. After fi rst spray, inspect 

foliage with a hand lens to determine if 

the spray was effective or if a second spray 

is necessary. 





Hosts 

• Eastern white pine (Pinus strobus) 

• Other fi ve-needled pines 

Alternate Hosts 

• Currants 

• Gooseberry 

• Other Ribes species 




On Pine 

Year-round: 

• Chlorotic needles, stunted growth 

• Chlorotic, dead, or dying tree tops or 

branches 

• Yellow-bordered cankers on trunk or 

3- to 4-year-old branches; oozing resin or 

rodent feeding may be apparent 

• Resin fl ow on main trunk that hardens to 

white, orange, or brown 

Early spring: 

• Orange-yellow blisters breaking through 

cankered bark to release spores 

Late spring to early summer: 

• Sticky, yellow fl uid produced from 

yellow-brown blister on canker; fl uid 

will blacken after a short time 

Fall: 

• Patches of yellow or brown bark on 

young growth; area may be swollen but 

progresses to spindle-shaped canker by 

second year of infection 

On Ribes Species 

Summer: 

• Orange spores on underside of leaf; upper 

surface may have yellow, diffused spots 

Late summer to early fall: 

• Brown, hairlike projections on underside 

of leaf 

Figure 1. Flagging as a result of cankers girdling 

a branch. Courtesy of John W. Schwandt, USDA 



• Fungal root rots 

• Pales weevil 

• Pine root collar weevil 


White pine blister rust is the only stem rust 

of fi ve-needled pines. It requires an alternate 

host (Ribes species) for new infections 

to occur on pine. 

Early detection can be diffi cult due to 

very subtle symptoms during the fi rst year of 

infection. First, a small, yellow or red spot 

appears on the needle at the site of infection 

and eventually the needle may die as the 

fungus grows into the bark tissue. The newly 

infected bark tissue will become brown with 

a yellow border surrounding a section of dead 

needles. The stem may be slightly swollen. 

During the second year of infection 

and beyond, long, elliptical-shaped cankers 

develop on branches and the main trunk as 

the fungus advances. Cankers can eventually 

girdle the branch, which results in 

“fl agging” (Fig. 1), or they can girdle the 

trunk, killing all growth above the canker. 

Yellowish-green bark tissue may be visible 

around the canker. Blisters (0.25 inch, 

3 mm across), spores, and resin fl ow all arise 

from the canker area. Rodent feeding may 

be evident around this area as well. 


The life cycle of white pine blister rust may 

take 3–6 years to complete. It begins in late 

summer or early autumn when basidiospores 

from the alternate host (Ribes species) 

are wind and rain dispersed, entering the 

pine needle through the stomata (Fig. 2). 

Basidiospores may be carried in wind 

currents for up to a mile. 

Figure 2. Yellow/red infection site on a needle. 

Courtesy of USDA Forest Service Ogden 

Archive, Bugwood.org (#1467424) 

WHITE PINE 

BLISTER RUST 

Cronartium ribicola 

J. C. Fisch 

Elliptical cankers covered 

with yellow blisters that 

release infectious spores to 

be carried away by the wind. 


PDA 

WHITE PINE BLISTER RUST .................................................................................................................................................................................................................................................................... ..................................................................... 71 99

Figure 3. Elliptical cankers 

formed after the fi rst year of 

infection. Courtesy of Joseph 

O’Brien, USDA Forest Service, 


Figure 4. Mass of powdery, 

yellow spores covering the 

surface of the canker (spring). 


Figure 5. Sticky, yellow liquid 

exuded by a canker just before 

hardening off. Courtesy of 

Joseph O’Brien, USDA 


(#5042098) 

The fungus grows into the bark tissue 

at a rate of 5–6 inches (12.5–15.0 cm) per 

year and begins to form cankers after the 

fi rst year of infection (Fig. 3). In spring, 3–4 

years after the initial infection, pale yellow 

or cream-colored blisters (aecia) rupture 

through the bark of active cankers (Fig. 

4). They release powdery, yellow spores 

(aeciospores) that are carried in the wind 

over long distances to the alternate host 

and cause infection. The aeciospores can 

only infect Ribes species. After the spores 

are released, the cankered area on the pine 

remains swollen and roughened. In summer, 

the sticky, yellow fl uid that exudes from the 

site hardens and leaves small, brown-rustcolored 

scars (Fig. 5). 

On the alternate host, the aeciospores 

enter the stomata of the leaf during wet 

weather. Diffused, yellow spots become 

visible on the upper leaf surface soon after 

infection occurs. Within a few weeks, pustules 

form on the leaf underside and release 

spores that repeatedly infect the same plant 

or other Ribes in the vicinity (Fig. 6). This 

repeating stage serves to increase the levels 

of inoculum. In late summer, small, brown, 

hairlike structures appear on the underside 

of the leaf. Eventually, basidiospores 

are produced and wind dispersed back to 

susceptible pines in the vicinity. 

Figure 6. Underside of Ribes leaf with 

fruiting bodies. Courtesy of Robert L. 

Anderson, USDA Forest Service, 



Strategies 


• Northeastern Pennsylvania is especially 

prone to white pine blister rust. 

• Do not plant white pine in low-lying areas 

where cool, moist air is likely to settle. 

• Do not plant white pine species if there 

is an abundance of alternate hosts 

(Ribes spp.) in the surrounding area. 

• Consider removing alternate host 

material from within 1,000 feet of 

white pine plantings. 

Preseason 

• Tag trees suspected to have white pine 

blister rust to check for blisters in 

mid-April. 


• Scout trees for blisters in mid-April. 

— Randomly select at least 50 trees that 

are 5–10 years of age. 

— Look for yellow or orange blisters on 

branches and trunks. 

— Also scout tagged trees. 

• If 10 percent or more of sampled trees are 

infected: 

— Prune/remove cankers from trees before 

infection reaches trunk. 

— Control alternate host before August. 

• Remove and destroy trees with trunk 

infections. 

• Inspect trees throughout the year for 

cankers. 




Biological 


at this time. 

Mechanical 

• Prune and destroy all branches with 

cankers. 

• Remove and destroy all alternate host 

plants in and around plantation within a 

minimum of 1,000 feet. 

Biorational 


at this time. 

Chemical 

• Apply an appropriate fungicide in late 

summer to protect pines from infection 

from basidiospores released by an alternate 

host. 

• Apply an appropriate herbicide to control 

the alternate host. 


• Inspect plants/nursery stock; buy from a 

reputable company. 

SHOOT AND BRANCH INJURY ............................................................................................................................................................................................................................................................ 100

Hosts 

In some areas, the pest is called the Sitka 

spruce weevil or Engelmann spruce weevil 

because of preference for these hosts: 

• Spruce 

• Pine 


• Fir (occasionally) 


• Moderate–high (depending on stand age 

and species of host) 


Spring 

• Adult weevils resting on terminal bud 

or on bark of previous year’s terminal; 

mating pairs are common; may drop to 

ground when approached 

• Pin-sized holes in the bark of terminal 

leader; clear droplets of sap oozing 

from holes; best observed on dry, sunny 

mornings on eastern white pine 

• Oval, cream-colored eggs in depression 

under feeding site 

Summer 

• Spongy, softened area under bark of last 

year’s terminal (bark may be discolored in 

this area) 

• White, legless grubs with medium brown 

head feeding in cambium tissue under 

bark of leader; multiple larvae usually 

present, often feeding in a ring around terminal; 

tissue above larvae reddish brown 

• By July, new terminal growth wilting 

or drooping, forming characteristic 

shepherd’s crook; Douglas-fi r generally 

wilts fi rst, followed by eastern white 

pine; spruce damage usually last to be 

noticeable 

• Terminal dead down to second or third 

whorl of branches 

• Oval areas fi lled with shredded wood 

located under bark or inside the dying terminal; 

whitish pupae under wood shreds 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

• In late summer, round adult emergence 

holes about 2 ⁄25 inch (2 mm) in diameter 

in bark on trunk below dead terminal 


• Tends to attack a tree in successive years, 

resulting in deformed or forked trees 


• Frost injury 

• Bird damage 



• Zimmerman pine moth 


White pine weevil adults have long snouts 

typical of all weevils. Elbowed antenna 

with terminal knobs is another identifying 

characteristic for weevils. The body of 

white pine weevil is covered with broad, 

scalelike hairs that vary from medium brown 

to almost black. Patches of white and gold 

scales are noticeable on their wing coverings. 

In appearance, they are almost identical to 

eastern pine weevil, differing only in size. 

White pine weevil is the smaller of the two 

pests, measuring about ¼ inch (6–7 mm) 

long; males are slightly smaller than females. 

The larvae are white, legless, C-shaped grubs 

with a medium brown head and several long, 

silken hairs on the body. 

It is diffi cult to see weevils on many of 

the host trees because of their color, which 

blends well with most conifer bark. Adults 

are easiest to fi nd on the smooth bark of 

eastern white pine terminals. White pine 

weevil can also be diagnosed by the damage 

and chip cocoons under the bark of the dead 

terminal. No other weevil species feeds and 

pupates in the terminal of healthy trees. 

White pine weevil tends to attack a tree 

in successive years, resulting in deformed 

or forked trees. However, young trees may 

be affected if found next to fi elds with high 

pressure. Scouting reports in Pennsylvania 

have shown fi elds infested at all ages. 


Adults emerge. 

WHITE PINE 

WEEVIL 

Pissodes strobi (Peck) 

Typical “shepherd’s crook” 

symptomatic of white pine 

weevil infestation. Courtesy 

of PDA 

WHITE PINE WEEVIL .............................................................................................................................................................................................................................................................................. ................................................................... 101 71

Figure 1. Adult weevils. 

Courtesy of Dave Powell, USDA 


(#1207040) 

Figure 2. Adult weevil (circled) 

on Colorado blue spruce. 


PDA 

Figure 3. Sap droplets seeping 

from feeding or egg holes. 


PDA 


White pine weevils overwinter as adults in 

the ground litter, or duff, under trees (Fig. 

1). As the weather warms in mid-March 

or April, adults emerge from overwintering 

areas. Since attacks commonly occur 

in subsequent years on the same tree, the 

overwintering site is frequently under the 

same tree that will be attacked in the spring. 

When weevils become active, they crawl up 

the trunk until they reach a healthy leader 

(Fig. 2). On warm days, they often fl y at 

canopy level to other trees. For 2–3 weeks, 

both males and females feed just below the 

terminal bud by chewing through the bark 

to make a pin-sized hole. With their strong 

mouthparts, they feed by scooping out the 

cambial layer surrounding this hole. 

Overwintering females were generally 

mated the previous fall and many retain 

viable sperm through the winter. However, 

on warm spring days, mating pairs are 

frequently found on terminal shoots. When 

eggs are mature, the female chews a fresh 

hole in the bark and scoops out cambial 

tissue to create a small egg niche. She usually 

deposits a single egg and seals the hole with 

dark brown frass before moving to another 

nearby site and repeating the process. Egg 

sites will have a droplet of clear sap (Fig. 3). 

Female white pine weevils deposit an average 

of 100 eggs, which hatch 6–14 days later. 

The whitish, oblong eggs measure 1 ⁄25 inches 

(1 mm) long (Figs. 4 and 5). 

Figure 4. Two weevil eggs sharing 

an egg niche. Courtesy of Sandy 

Gardosik, PDA 

Figure 5. Pale, 1-millimeter, 

oblong eggs under the bark and 

cambium layers. Courtesy of 


When the larva emerges from the egg, 

it tunnels upward toward the bud before 

turning and moving downward (Fig. 6). 

This feeding is generally restricted to the 

previous year’s growth. It is common to 

have several larvae feeding in a ring around 

the terminal (Fig. 7). This effectively cuts 

off all fl ow of nutrients and moisture to the 

new growth, causing it to wilt and die. If the 

number of larvae is too high, competition 

will result in death of some larvae; if too 

low, the larvae may drown in pitch. When 

larvae drown in pitch, the terminal will 

survive but be deformed. Larvae from eggs 

deposited later in the season frequently do 

not survive because of lack of food. 

Figure 6. Larvae feeding on vascular tissue. 


By midsummer, mature larvae are 2 ⁄5 inch 

(1 cm) long. Each larva creates a pupal cell 

under the bark or in the pith of thinner terminals. 

The larvae use wood strands to form 

characteristic chip cocoons before they pupate 

(Fig. 8). Pupation takes several weeks, 

and the weevil may remain in this cell for 

as long as 6 weeks (Fig. 9). When adults 

fi rst emerge, they are very pale and soft and 

referred to as callow adults. These callow 

adults can be found in the chip cocoon until 

their body has hardened, at which time they 

chew a round hole in the bark and emerge 

(Fig. 10). Adult emergence usually starts 

in mid-July and may continue into early 

September. Weevils will feed on branches of 

the host tree and mate before moving to the 

ground litter to overwinter. Most adults only 

overwinter one time, but reports of adults 

living for several years can be found in the 

literature. 


Figure 7. Larvae feeding around 

the terminal. Courtesy of Daniel 

Herms, The Ohio State University, 


Figure 8. Chip cocoon. Courtesy of 


Figure 9. Pupae maturing in chip 

cocoons. Courtesy of Sandy 

Gardosik, PDA 

Figure 10. Adult weevil emerging 

from bark. Courtesy of Rayanne D. 

Lehman, PDA 


Strategies 


• Remove any unmanaged trees that may 

act as pest reservoirs. 

• Preferred attack sites include open-grown 

trees in full sunlight. Terminals that are 

1 

⁄5 inch (5 mm) in diameter are preferred. 

Planting in partial shade may reduce 

attack by keeping the temperature lower, 

inhibiting weevil activity. 

Preseason 

• Train new tree leaders for last year’s 

damaged trees by using a healthy lateral 

branch from the previous season’s growth. 

• Late winter/early spring: Place modifi ed 

Tedder’s traps (see Appendix E) in the 

fi eld before adult weevils become active 

on a daily basis. Place traps in the row 

near trees that sustained weevil damage 

the year before (Fig. 11). Check traps 

regularly to monitor weevil activity (Fig. 

12). Traps must be baited with denatured 

alcohol and turpentine to be attractive to 

weevils. 

• Although trapping is the preferred detection 

method, do not overlook scouting on 

sunny days by examining leaders for adults 

or droplets of clear sap. Droplets will glisten 

in the sunlight (see Fig. 3 on p. 102). 

• Using binoculars is helpful for those short 

in stature. 

• When resin droplets are found, scrape 

away the sap to reveal if there is a circular 

feeding site. 

• Growing degree days: Adults generally 

become active from 7 to 58 GDDs. 

• Ground temperatures: After a 3-year 

monitoring project in Pennsylvania, it was 

determined that adult white pine weevils 

become active when ground temperatures 

are above 50°F. Insert a probe thermometer 

2 inches into the soil to collect soil 

temperature readings (Fig. 13). This must 

be monitored on the sunny side of the 

tree, in warmer fi elds, or on south-facing 

slopes fi rst. 

Figure 11. Tedder’s trap placed 

among previously damaged 

trees. Courtesy of Rayanne D. 

Lehman, PDA 

Figure 12. Adult weevils caught 

in a Tedder’s trap (plastic top 

has been removed). Courtesy of 


Figure 13. Soil thermometer. 


WHITE PINE WEEVIL ............................................................................................................................................................................................................................................................................. 103



established for this pest in Christmas 

trees. In forestry, 2–5 percent of trees 

damaged in the previous season is used as 

a treatment threshold. 

• Check leaders for continued feeding to 

determine the need for additional sprays. 

• When the injury becomes too severe to 

manage with pruning, treat the entire 

plantation. 




Biological 

• White pine weevil has several naturally 

occurring parasitoids, such as Lonchaeid 

fl ies and wasps, but they do not provide 

adequate control. 

• Birds may feed on larvae and pupae. 

• Rodents will consume overwintering 

adults. 

Mechanical 

• Prune damaged leaders as soon as wilting 

is detected but before adults emerge in 

mid-July. Leaders must be pruned down 

to healthy, green wood. 

• Damaged leaders should not be used to 

train new leaders. 

• Damaged tissue containing larvae or 

pupae should be removed from the fi eld 

and destroyed. Depending on the time 

of year and weather following pruning, 

adults may still be able to emerge from 

leaders dropped to the ground after 

pruning. 

Biorational 


at this time. 

Chemical 

• Chemical controls should begin as soon 

as the fi rst white pine weevil is trapped 

or feeding sites or adults are seen on 

terminals. 

• Spray applications must be made 

promptly. Delaying treatments results 

in poor control. 

• If a sharp drop in temperature is forecast 

following the fi rst emergence of the 

weevils, wait to apply sprays until the 

next set of weevils is found in traps. 

• Controls should only be applied to the 

top third of the tree. 


• Remove any damaged or highly susceptible 

trees surrounding the plantation 

before planting. 


STEM AND ROOT INJURY/ 

TREE MORTALITY 

Entire trees may be killed by the diseases 

and insects on pages 107–121. Some of 

these may be associated with other pests 

that initially weaken a tree but are not 

capable of killing it. The combination 

of pests can result in rapid death of some 

conifers. 

STEM AND ROOT INJURY/TREE MORTALITY ...................................................................................................................................................................................................................................... 106

Hosts 

• Most species of Christmas trees 

• Approximately 700 different species of 

woody plants and some herbaceous plants 




• Reduced terminal growth 

• Yellowing and eventual browning of 

needles 

• Whitish resin at the base of tree 

• Creamy-white fans of fungus between 

bark and wood at the root collar 

• Brown to black fungal rhizomes resembling 

shoestrings beneath the bark, on 

the roots, and in the soil 

• Groups of tan-colored mushrooms near 

decaying wood in autumn 

• Death of young trees, often in groups 


• Other root and canker diseases 


• Wood borers 


• Drought 

• Wet feet 


Many known species of Armillaria exist in 

North America and are not easily distinguishable 

from one another. The most common 

and pathogenic species to conifers are 

Armillaria mellea (Vahl:Fr.) P. Kumm. and 

A. ostoyae (Romagnosi) Herink. Other 

common names for Armillaria include 

shoestring root rot, oak fungus, honey 

mushroom, and honey agaric. 

Conifers frequently show symptoms 

and signs of Armillaria infection at different 

rates. Trees under stress, such as during the 

heat of summer or drought, are more likely 

to become infected and show symptoms. 

Young trees may show symptoms and die 

very rapidly compared to older trees, which 

may show symptoms for years before dying. 

Characteristic signs of Armillaria are 

visible beneath the lower trunk bark and in 

the soil. The fungus produces long, black, 

stringlike strands called rhizomorphs, which 

can easily be mistaken for small roots. They 

are found between the bark and wood, on 

the surface of roots, and in the adjacent soil 

(Fig. 1). In addition, creamy-white, paperthick, 

fan-shaped sheets of the fungus can 

be seen when bark is removed at the tree 

base (Fig. 2) or when it is exposed in the 

main roots (Fig. 3). 

Figure 1. Black, stringlike strands, or rhizomorphs, 

between the bark and wood of the 

lower trunk and in the soil. Courtesy of Joseph 

O’Brien, USDA Forest Service, Bugwood.org 

(#5047087) 

Figure 2. Creamy white, paper-thin, fan-shaped 

sheets of mycelium under the bark near the 

tree’s base. Courtesy of Minnesota Department 

of Natural Resources Archive, Bugwood.org 

(#4214008) 

Figure 3. White fungal sheets found under 

the epidermis of the tree’s roots. Courtesy of 

Edward L. Barnard, Florida Department of 

Agriculture and Consumer Services, Bugwood 

.org (#4822096) 

ARMILLARIA 

ROOT ROT 

Armillaria spp. 

Needle yellowing and 

browning, a symptom of 

Armillaria. Courtesy of Joseph 

O’Brien, USDA Forest Service, 


ARMILLARIA ROOT ROT ...................................................................................................................................................................................................................................................................... ................................................................... 105 107

Figure 4. Mushrooms produced 

by some Armillaria species near 

the trunk of infected trees. 

Courtesy of Joseph O’Brien, 


.org (#5047089) 

Figure 5. Tree death as a result 

of Armillaria infection. 

Courtesy of Steven Katovich, 


.org (#1479015) 

Diseased wood fi rst looks water soaked 

and light brown in color. It becomes yellowish 

white, spongy, and stringy over time. 

Prolonged decay can also cause vertical 

cracks in the root collar. Armillaria has a 

very strong mushroom odor and some species 

produce clusters of yellowish-brown 

mushrooms (associated with the rhizomorphs) 

near the decaying wood after a 

period of rain in the fall. 


Some species of Armillaria will produce 

mushrooms near the trunk of infected trees 

(Fig. 4). Though new infections can result 

from airborne spores released by these mushrooms, 

the most common means of disease 

spread is by underground growth of the 

rhizomorphs originating from an infected 

tree. Rhizomorph structures can survive for 

many years on dead or dying tree roots and 

stumps and spread through the soil up to 

60 feet from the point of origin. As healthy 

tree roots grow through the soil, they come 

into contact with the rhizomorphs and 

mycelium. By secreting an enzyme that 

breaks down cell walls, the rhizomorphs and 

mycelium adhere to the healthy tree roots 

and penetrate into them. The fungus infection 

causes the loss of the tree’s fi ne feeder 

roots and results in insuffi cient water and 

nutrient transport to the trees, which leads 

to tree decline and death (Fig. 5). If the soil 

temperature reaches 79°F (26°C), Armillaria 

growth can be inhibited. 


Strategies 


• Do not plant in a recently cleared 

hardwood stand that had a previous 

problem with Armillaria. 

• Avoid areas surrounded by oak forest. 

• Avoid areas that may cause extreme 

tree stress such as drought or excessive 

moisture. 

Preseason 

• No scouting schedule or technique is 

available for this disease. However, when 

scouting for other insect pests or diseases, 

watch for declining trees that may be 

infected with Armillaria. 


• Test soil and maintain proper nutrient 

balance. 

• Water when there is a drought. 

• Control other pests affecting trees during 

the season to maintain plant vigor. 




Biological 


at this time. 

Mechanical 

• Dig up and remove diseased trees, tree 

stumps, roots, pruning waste, and all 

infected wood and burn it on site. 

• If only a few roots are infected, remove 

the infected soil from midspring through 

late fall to expose the root collar and 

buttress roots to air and sun. This allows 

them to dry out. Before the fi rst heavy 

frost, replace the removed soil with 

Armillaria-free soil. 

Biorational 


at this time. 

Chemical 

• Fungicides are not recommended for 

treatment for this disease. 

• Soil fumigants have been used with 

limited success after diseased material 

is removed from the soil. However, soil 

fumigants are highly toxic and should 

only be applied by a licensed pesticide 

professional. 




Hosts 

• Pines 


• Low (trees in fi eld) to severe 

(balled-and-burlapped eastern white pine) 


• Needles turn yellow to red 

• Dying or dead parts of tree or whole dead 

trees 

• Galleries (tunnels) under loose bark 

• Dry, reddish-brown boring dust in bark 

crevices and around gallery openings on 

main trunk 

• White larvae or dark brown adult beetles 

in the galleries 



• Pine wilt disease 

• Armillaria root rot 


Bark beetles are small, cylindrical beetles 

about the size of a grain of rice. They are 

usually dark brown to black, but newly 

emerged adults are lighter in color. It may 

be diffi cult to see the head on some species. 

The chestnut brown bark beetle (Pityogenes 

hopkinsi) is about 1 ⁄ 16 inch (1.6 mm) long 

and is limited to eastern white pine. Ips 

adults are slightly larger, ranging from 1 ⁄ 8 

to 3 ⁄ 16 inch (3.18 to 4.76 mm) in length. 

Ips beetles have distinct spines on their 

posterior end. All species of pine are hosts 

for one or more species of Ips beetle. The 

Ips beetles are often referred to as engraver 

beetles because of the types of galleries they 

create under the bark. 

Larvae of these beetles are legless, 

creamy white, and have light brown head 

capsules. They may reach 3 ⁄ 16 (4.76 mm) 

inch long when fully grown, depending on 

the species. 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 


Overwintered males of both engraver 

(Ips) and bark (Pityogenes) beetles initiate 

an attack on a tree weakened by disease, 

environmental conditions, or other pests 

(Fig. 1). Each male chews a small gallery 

under the bark and deploys an aggregation 

pheromone, which attracts both sexes of 

the same species to the target tree (Fig. 2). 

In addition to bringing females for mating, 

the pheromone attracts more males to begin 

additional galleries. As a result, the beetles 

Figure 1. Adult bark beetle. Courtesy of Jim 

Stimmel, PDA, Bugwood.org (#2120093) 

Figure 2. Gallery of the bark beetle, Pityogenes 

hopkinsi. Courtesy of Rayanne D. Lehman, PDA 


BARK AND 

ENGRAVER 

BEETLES 

Ips spp. and 

Pityogenes hopkinsi 

Swaine 

Damage from a bark beetle 

attack. Courtesy of PDA 

BARK AND ENGRAVER BEETLES ......................................................................................................................................................................................................................................................... ................................................................... 107 109

Figure 3. Bark beetle boring 

dust on pine trunk (symptom 

of beetle attack). Courtesy 

of Jeffrey Eickwort, Florida 

Department of Agriculture and 

Consumer Services, Bugwood 

.org (#5179069) 

Figure 4. Engraver beetle (Ips 

spp.) larva. Courtesy of G. Keith 

Douce, University of Georgia, 


Figure 5. Ips spp. callow adult, 

larva, and pupa (from left 

to right). Courtesy of Roger 

Anderson, Duke University, 


generally attack a single tree en masse (Fig. 

3). As the beetles enter the gallery, they 

introduce fungal spores, which quickly 

germinate. The fungus serves two purposes: 

(1) food for larvae, and (2) more important, 

it blocks resin fl ow, the tree’s main 

defense against bark beetle establishment. 

Many adults die in the attack, but the tree’s 

defense is eventually overcome. 

Females that enter the male’s initial gallery, 

or nuptial chamber, mate with the male 

and then continue chewing an egg gallery. 

Individual eggs are laid in niches along this 

gallery. When larvae emerge, they each continue 

a gallery radiating out from their egg 

niche (Fig. 4). Engraver beetles may have 

Y- or H-shaped galleries in the inner bark. 

Bark beetles, on the other hand, generally 

have a star-shaped gallery. Larvae pupate 

in the galleries (Fig. 5) and adults emerge 

through the bark to continue their life cycle 

on a different tree. 

Engraver beetles can complete a generation 

in 21–40 days under optimum conditions. 

How many generations are completed 

each year is not known. Bark beetles appear 

to have two main fl ight periods—one in 

spring and one later in summer. 


Strategies 


• Select a site for optimal pine growth. 

• Maintain proper spacing when planting. 

• Remove potential breeding material 

(e.g., mature pines damaged by disease, 

insects, wind, or other harmful causes) 

from the perimeter of the block. 

Preseason 

• Remove any dead or dying pines that may 

act as reservoirs of bark beetles. 

• Chip piles or tree residue can also be 

reservoirs. 


• Pityogenes hopkinsi will attack eastern 

white pine under stress during and after 

digging (Fig. 6). Maintain adequate 

moisture to roots and replant as soon as 

possible to lessen stress. 

• Threshold level: Regulatory concern 

exists for dug trees. Consult your local 

state plant inspector for assistance. 

• Maintain tree vigor by fertilizing and 

irrigating as required. 




Biological 

• Insect parasites and predators and fungal 

diseases offer some control. Woodpeckers 

will feed on engraver (Ips) beetles, 

especially during the winter. 

Mechanical 

• Remove and burn, bury, chip, or debark 

infested parts of trees or dead trees to get 

rid of the insects. 

Biorational 


at this time. 

Chemical 

• Chemical control is only necessary for 

high-valued trees in recreational or 

residential settings. Stressed balledand-burlapped 

white pine may require 

chemical applications during beetle 

fl ight periods. 

Figure 6. Balled-and-burlapped trees awaiting 

delivery—prime targets for bark beetles. 

Courtesy of Rayanne D. Lehman, PDA 


Hosts 

• Most Christmas tree species 

• True fi rs, Douglas-fi r, spruce, and eastern 

white pine highly susceptible 


• High 


• Reduced or stunted growth 

• Chlorotic or red-brown needles 

• Needle loss 

• Root decay 

• Bleeding basal cankers 

• Death of the tree 


• Armillaria root rot 

• Procera root rot 



Phytophthora identifi cation requires laboratory 

analysis, but some symptoms in the fi eld 

should make the grower suspect this disease. 

Trees that do not thrive after planting or 

quickly develop reddish-brown needles 

and exhibit dieback should be checked for 

Phytophthora. Examine roots for symptoms 

of decay and absence of an extensive feeder 

root system (Fig. 1). Low-lying areas with 

poor water drainage are especially prone to 

root rot diseases such as Phytophthora (Fig. 2). 

Figure 1. Roots decay and trunk base 

may discolor from Phytophthora infection. 



Phytophthora is a soilborne water mold that 

can spread from an infested fi eld to a new 

fi eld through the movement of water in the 

soil or on the surface. New infections can 

occur when the temperatures exceed 59°F 

(15°C). Resting spores (chlamydospores 

and oospores) that are capable of surviving 

for many years in the soil or plant are 

formed during cold and/or dry periods. When 

required temperature and moisture conditions 

are present, these resting spores will 

germinate and form another type of sporeproducing 

structure called a sporangia. When 

mature, numerous motile, infectious spores, 

or zoospores, are released. Zoospores swim 

for up to an hour through the soil water and 

are attracted to the plant roots by chemicals 

that are produced during growth. When they 

come in contact with susceptible tissue, they 

germinate and penetrate into the roots, form 

mycelium, and cause infection. Zoospores are 

spread farther distances from an infested fi eld 

to a new fi eld through the movement of fl owing 

surface water. It should be assumed that 

any plant, soil, or water that is transported 

from an infested fi eld is contaminated with 

some type of Phytophthora spore. 

Figure 2. Phytophthora infection often follows 

the slope of a hill. Courtesy of Tracey Olson, 

PDA 

PHYTOPHTHORA 

ROOT ROT 

Phytophthora spp. 

Severe tree browning, a 

symptom of Phytophthora 

root rot. Courtesy of Tracey 

Olson, PDA 

PHYTOPHTHORA ROOT ROT .............................................................................................................................................................................................................................................................. ................................................................... 107 111

Figure 3. Browning and loss 

of infected tree feeder roots. 

Courtesy of Edward L. Barnard, 

Florida Department of Agriculture 

and Consumer Services, 


Figure 4. Sudden wilting and 

browning of infected tree. 


As the mycelium continues to develop 

inside of the roots, the roots will die and 

turn brown (Fig. 3). The fungus will spread 

from the outer roots toward the larger roots, 

the root crown, and eventually the stem. 

The conductive tissue of the plants will decay 

and prevent fl ow of water and nutrients 

to the upper portion of the tree. Needles 

will fi rst turn chlorotic and then a reddish 

brown, while branches wilt (Fig. 4). This 

infection will lead to death of the tree. 


Strategies 


• Buy and plant healthy seedlings from a 

registered grower. 

• Do not plant in a fi eld known to be 

infected with Phytophthora. 

• Monitor seedlings before planting. Look 

for plants that show reddish-brown roots 

or other symptoms of root rot and do not 

plant these in the fi eld. 

• Have suspicious plants analyzed for 

Phytophthora by a diagnostic lab. 

• Avoid planting Fraser fi r in areas that 

retain considerable moisture. 

• Some growers in Pennsylvania have tried 

mounding the soil in the rows before 

planting as a preventative strategy, 

though no research has been done on 

the effectiveness of this technique. 

Preseason 

• Do not overfertilize or overwater trees. 


• Monitor fi elds for symptomatic trees. If a 

tree is suspected to be infected, remove 

the tree and the root ball from the fi eld 

and burn them, unless the tree is to be 

tested for the pathogen. 

• To receive confi rmation that the pathogen 

is Phytophthora spp., send sample to 

an extension laboratory or state regulatory 

laboratory. 

• Trees that are in close proximity to known 

Phytophthora-infected trees should be 

marketed only as cut Christmas trees as 

soon as possible. 




Biological 


at this time. 

Mechanical 


at this time. 

Biorational 


at this time. 

Chemical 


at this time. 


• Replanting with susceptible hosts in 

known Phytophthora-infected fi elds is not 

recommended. 

• Most conifers grown in Pennsylvania 

are susceptible to Phytophthora spp. to 

some degree. Tolerance to Phytophthora 

has been shown with some species 

of Abies, most notably Turkish (A. 

bournmuelleriana) and Momi (A. fi rma). 

• Some success has occurred with using 

grafted Fraser fi rs to Turkish and Momi 

fi r rootstock in Phytophthora-infected 

sites, though sources for these plants are 

limited. 


Hosts 

• Scotch, Austrian, and red pine common 

breeding sites; scotch pine is the least 

tolerant of attack; eastern white pine generally 

resistant due to excessive sap fl ow 

• Open-grown, healthy trees from 2 to 20 

feet (0.61–6.1 m) tall preferred; trees one 

inch or less in diameter at root collar are 

not suitable hosts 

• Adults readily feed on bark of eastern 

white, Scotch, and red pine 




• First external symptoms: trees loose in 

soil, leaning, or dying (entire tree may be 

yellowed; infested trees may live several 

years before turning brown and dying) 

• Presence of blackened, pitch-soaked soil 

and/or white resin around the root collar 

• Root collar at or below soil line restricted 

or entire trunk broken off 

• Active infestations will have one or more 

white, legless larvae or pupae in the soil or 

under bark 


• Root rot diseases 

• Mechanical damage 

• Rodent damage 


The legless, C-shaped larvae have brown 

heads and whitish bodies. They closely 

resemble larvae of other weevil species that 

also attack pines. Mature larvae are ½– 2 ⁄3 

inch (14–17 mm) in length. A good fi eld 

identifi cation tip for this pest is the location 

of the larvae. Pine root collar immatures 

are only found in the root collar area below 

ground level. To locate larvae, remove soil 

down to the fi rst root fl are. They may be 

in the cambial layer of the host or in the 

surrounding pitch-soaked soil. Occasionally, 

pitch tubes extending from the root collar 

into the soil will be found. 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

Pine root collar weevils never pupate in 

a typical chip cocoon, unlike other conifer 

weevil pests. Instead, they create their pupal 

cell surrounded by soil or pupate in their 

gallery in the bark of the root collar area. 

Adults are typical weevils with elongated 

snouts and elbowed antennae. They 

are dark brown with small patches of fi ne, 

pale hairs on the wing covers. The adults 

resemble pales weevil adults but are slightly 

larger, ranging from 1 ⁄3 to ½ inch long (10.0– 

12.5 mm) (Fig. 1). Consult a plant inspector 

or entomologist to confi rm identifi cation. 

Figure 1. Adult pine root collar weevil (left); 

pales weevil (right). Courtesy of Jim Stimmel, 

PDA 


Only adults are active above the soil line, 

where they will feed on tender bark of pine 

twigs. They are nocturnal feeders and prefer 

to feed on the underside of lower branches. 

Adults also move from tree to tree at night 

but crawl on the ground since they are weak 

fl iers. They must be careful of their movements 

during the summer because a soil 

surface temperature greater than 104°F will 

kill them. Adults also feed with the larvae 

on the inner bark of the root crown. 

The life cycle of pine root collar weevil 

is completed in two seasons. In early May, 


PINE ROOT 

COLLAR WEEVIL 

Hylobius radicis 

Buchanan 

Dying tree with pine root 

collar weevil infestation. 


PDA 

PINE ROOT COLLAR WEEVIL ............................................................................................................................................................................................................................................................... ................................................................... 107 113

Figure 2. Egg laying and larval 

feeding within the root collar 

just below the soil. Courtesy of 


Figure 3. Larva feeding on the 

root collar. Courtesy of James B. 

Hanson, USDA Forest Service, 


Figure 4. Excessive pitch in the 

soil surrounding the root collar 

resulting from larval feeding. 


PDA 

Figure 5. Pupa in soil near the 

root collar. Courtesy of Sandy 

Gardosik, PDA 

young overwintering adults emerge. Mating 

and egg laying occur at the base of the tree 

during the day and egg laying may continue 

into September. Eggs are deposited 

in feeding wounds made by an adult in the 

inner bark of the root collar (Fig. 2) or a 

few centimeters away from a root in the soil. 

Two weeks later, the eggs hatch and young 

larvae start feeding on the inner bark of the 

root collar area (Fig. 3). This causes pitch 

(or sap) to fl ow into the surrounding soil, 

eventually reaching 15–20 inches (40–50 

cm) from the trunk (Fig. 4). Larvae create 

tubes in this pitch-soaked soil to drain excess 

pitch away from the wound area. They 

may also use the pitch tubes for molting and 

pupating. 

The majority of larvae overwinter in the 

root collar as third and last instars. However, 

with the length of egg laying, early 

eggs may produce adults in the same season. 

Overwintered larvae become active the 

following spring, complete development, 

and pupate in late June to mid-August 

(Fig. 5). The new adults begin to emerge in 

late summer and feed on pine bark before 

overwintering in litter under the tree or 

under the bark. These adults emerge the following 

spring to mate and lay eggs and may 

overwinter a second winter. Second-year 

adults will emerge in the spring to continue 

feeding, mate, and lay eggs before dying in 

early summer. 

All life stages of this weevil can be found 

during the growing season. During the winter, 

only large larvae and possibly pupae and 

adults will be present. Vigorously growing 

trees respond to larval feeding injuries by 

producing massive amounts of resin under 

the bark at and near the site of injury (Fig. 

6). Affected trees continue to live an average 

of 2–4 years after the initial attack (Fig. 7). 

Figure 6. 

Infected root 

collar soaked 

with resin. 

Courtesy of 

Steven 

Katovich, 

USDA Forest 

Service, 

Bugwood.org 

(#1199001) 


Strategies 


• Establish fi elds at least one mile away from 

any known infestations. 

• Plant least resistant varieties such as 

Scotch pine in small, single-species 

blocks. 

• Interplant pines only with fi r, spruce, or 

deciduous trees; do not interplant Scotch 

pine with other pines. 

Preseason 

• Examine off-color trees for symptoms. 

Rocking trees by hand will reveal any 

loose in the soil. Follow up with an 

examination of the root collar area for 

other symptoms. 



established. 



Figure 7. Mixed pine stand with varying 

degrees of infestation; damage is worst along 

the block’s edge where trees are stressed 

from grass competition. Courtesy of Steven 

Katovich, USDA Forest Service, Bugwood.org 

(#1199002) 



Biological 

• Use of parasites is not practical because 

larvae are below the soil and usually 

protected by pine pitch. 

Mechanical 

• To increase soil temperature and therefore 

decrease existing populations and lessen 

the risk of infestations, butt-prune trees a 

minimum of 12 inches (30.48 cm) high. 

• Remove leaf litter from under the tree, 

allowing the sun to heat and dry the soil. 

• Remove 1–2 inches (2.54–5.08 cm) of soil 

away from the trunk base, aiding in creating 

an undesirable environment for adult 

activity and egg laying. 

Biorational 


at this time. 

Chemical 

• Insecticide applications are only effective 

against adults when they are active at the 

base of the host tree. 

— Make an application in mid-May to 

control adults before egg laying. 

— Make a second application in mid- 

August to mid-September to kill newly 

emerging adults. 

— Both applications are required for 

successful control. 

• Apply only registered products for your 

state. Follow the directions on the label 

thoroughly. 

PINE ROOT COLLAR WEEVIL ................................................................................................................................................................................................................................................................ 115

PINE WILT 

DISEASE 

Caused by pinewood 

nematode, 

Bursaphelenchus 

xylophilus 

(Steiner and Buhrer) 

(Nickle) 

Dead tree with 

reddish-brown needles still 

attached. Courtesy of 

A. Steven Munson, USDA 


(#1470133) 

Host 

• Pines, particularly Scotch pine 




• Considerable decrease of resin fl ow from 

wounds 

• Needles turn yellow then reddish brown 

through growing season (wilting) 

• Needles may remain attached for several 

months following sudden death of tree 

• Wood of affected trees dries out and 

completely lacks resin 


• Wood borers and bark beetles 

• Diplodia tip blight 

• Atropellis canker 


This pest can only be identifi ed using a 

microscope. Check dying trees for symptoms 

and, if pine wilt disease is suspected, have 

samples examined by a pest specialist to 

determine if nematodes are present in the 

wood. 


Pinewood nematodes spread from infected 

pines to healthy or stressed pines in the 

spring through contaminated pine sawyer 

beetles (Fig. 1). These long-horned beetles 

acquire the nematode from infested trees 

during their development within the tree. 

When new adult beetles emerge in spring, 

they transfer nematodes to healthy trees 

through feeding or to diseased trees during 

egg-laying activities. 

Figure 1. Pine sawyer beetle is a vector of 

pinewood nematode. Courtesy of L. D. Dwinell, 


The nematodes have two phases. The 

propagative phase occurs in the sapwood of 

infested trees and includes six stages from 

egg to adult. A single generation can be 

completed in as few as 5 days under ideal 

conditions. This enables the nematodes to 

rapidly develop extremely high populations 

(Fig. 2). As they feed in the resin ducts and 

cambial tissues, the tree’s water-conducting 

system fails, causing rapid wilt under dry 

conditions (Fig. 3). 

Figure 2. Pinewood nematode feeding on resin 

ducts in pines. Courtesy of USDA Forest Service 

Rocky Mountain Region Archive, Bugwood.org 

(#1442033) 

Figure 3. Severe wilt caused by a failure of the 

water-conducting system from nematode feeding. 

Courtesy of USDA Forest Service North 

Central Research Station Archive, USDA Forest 


STEM AND ROOT INJURY/TREE MORTALITY ... 

...................................................................................................................................................................................................................................... 116

The second phase of the nematode life 

cycle is the dispersal phase. This only occurs 

in the late stage of tree infection when 

pine sawyer pupae are present. Under these 

conditions, immature nematodes develop 

into a nonfeeding stage that attaches to the 

pupal cell of the beetle (Fig. 4). When the 

adult beetles emerge, the nematodes will 

have contaminated their respiratory system 

and are carried with them to new host trees. 

Nematodes leave the beetle and enter the 

shoots of a new tree through the wounds 

created by beetle-feeding activity. Within 

48 hours of introduction to a new, healthy 

host, the nematodes have matured into 

reproducing adults. 

Pinewood nematodes feed on epithelial 

cells and resin ducts of healthy trees. They 

also may feed on blue-stain fungi in dead 

and dying trees to sustain or build population 

levels (Fig. 5). The fungus feeding 

cycle is the most common stage in North 

America. 

Figure 4. Nematodes entering the pine sawyer 

pupae will be transferred to a new host when 

the pine sawyer adults emerge. Courtesy of 

USDA Forest Service North Central Research 

Station Archive, Bugwood.org (#1406269) 

Figure 5. Pinewood nematodes 

feeding on blue stain fungi found in 

weak and dying trees. Courtesy of 

USDA Forest Service North Central 

Research Station Archive, Bugwood 

.org (#1406276) 


Strategies 


• Avoid planting on traditionally dry sites. 

• Do not plant susceptible pines in areas 

where mean summer temperatures are 

above 68°F. 

Preseason 


at this time. 


• Maintain tree vigor through periodic 

fertilization and irrigation during times of 

dry weather. 

• Any trees exhibiting symptoms should be 

analyzed by a diagnostic lab to determine 

presence of pine wilt nematode. 




Biological 


at this time. 

Mechanical 

• Remove and destroy (burn, bury, and/or 

chip) trees to help prevent spread of pest 

to nearby, healthy pines in the spring, 

before beetle emergence. 

Biorational 


at this time. 

Chemical 


at this time. 



PINE WILT DISEASE ............................................................................................................................................................................................................................................................................... 117

WHITE GRUBS 

(MAY, JUNE, 

AND JAPANESE 

BEETLES) 

Phyllophaga sp., 

Polyphylla sp., 

Popillia japonica 

Newman 

White grub damage to 

seedlings. Courtesy of 

Jim Stimmel, PDA 

Hosts 

• All conifers 


• Moderate–severe on seedlings 

Note: Balled-and-burlapped trees are 

regulated for the Midwest and western 

United States and parts of Canada, 

according to the U.S. Japanese Beetle 

Harmonization plan. For more information, 

refer to nationalplantboard.org/docs/ 

jbcolumn.pdf. 


• Seedlings and young conifer saplings 

discolored (reddish brown) in late summer 

to early fall, leading to eventual death 

• Lateral and taproots chewed off or girdled 

• Small holes, ¼– 1 ⁄3 inch (6–8 mm) in 

diameter, in soil surface 


• Drought stress 

• Phytophthora root rot 

• Pales or eastern pine weevil feeding 


Larvae of several species of scarab beetles, 

including the Japanese beetle, feed on roots 

of conifer seedlings and transplants. Collectively, 

the larvae are referred to as white 

grubs and are similar in appearance. Mature 

larvae are 1–2 inches (25–55 mm) in 

Figure 1. Adult Japanese beetle feeding 

on conifers (rare). Courtesy of Sandy 

Gardosik, PDA 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

length. These C-shaped, grayish-white grubs 

have tan heads and visible jointed legs. 

The posterior end of the grub is enlarged 

and frequently darkened. Most species have 

sparse yellow hairs on the body. All white 

grubs have a set of minute spines around the 

anus. The location of these spines, known 

as the raster pattern, is used to identify some 

species. 

Adults are dark brown to black beetles 

up to an inch (25 mm) in length. The common 

Japanese beetle adult is more colorful 

and familiar to anyone who gardens. Adult 

Japanese beetles have rarely been reported 

to feed on tender bark during a serious 

outbreak (Fig. 1). 


Scarab larvae, or white grubs, overwinter 

in the soil (Fig. 2). In spring, they move to 

the upper 4 inches of soil and resume feeding 

on the roots of almost every plant they 

encounter. When fully grown, these grubs 

create soil-encrusted cells before pupating 

near the soil surface. Adults begin to emerge 

in May and June. After feeding and mating, 

females deposit eggs just below the soil surface. 

As larvae emerge from the eggs, they 

immediately begin to burrow underground. 

Adequate soil moisture is required for survival, 

and drought periods will signifi cantly 

decrease the population due to desiccation 

of young larvae. 

Figure 2. June beetle larva. Courtesy of 

Steven Katovich, USDA Forest Service, 



STEM AND ROOT INJURY/TREE MORTALITY ... 

...................................................................................................................................................................................................................................... 118

Through the summer, the larvae feed 

in the top 4 inches of soil. They may travel 

considerable distances in search of plant 

roots on which to feed. Their strong mouthparts 

are capable of cutting off taproots of 

smaller seedlings. When grubs feed on conifer 

seedlings, the roots are consumed and 

the tree may be girdled just under the soil 

surface (Fig. 3). In beds of young seedlings, 

the plants may appear to be pulled deeper 

into the soil as the grubs feed. From time 

to time, the grubs emerge to the surface to 

deposit feces outside their burrow. In this 

process, they leave behind round holes in 

the surface that resemble puncture holes 

from a pencil. 

As soil temperatures decrease in fall, 

grubs move lower in the soil. They burrow 

8–10 inches (20.32–25.40 cm) below the 

soil surface to remain below the frost line 

overwinter. Japanese beetles only require 1 

year to complete a life cycle, but members 

of the genera Phyllophaga and Polyphylla 

require up to 4 years. 

Damage in seedling beds can occur as 

early as the fi rst year after germination. 

However, in transplants, the most severe 

white grub damage generally occurs 2–3 

years after fi eld planting. This is when the 

grubs are mature and capable of severing 

the taproots. Plantings that were previously 

in pasture or grass are the most susceptible 

and, generally, the grubs were present before 

the trees were planted. 


Strategies 


• Corn fi elds are known reservoirs for 

Japanese beetles. Conifer fi elds planted 

adjacent to these could be more susceptible 

to grub damage. 

• Evaluate the present white grub population 

in early fall or spring before planting. 

• Dig a 10-foot-long (3.05 m) furrow. If 

one grub per 10 feet of furrow is found, 

consider treating before planting the 

following year. 

• Dig several square-foot holes and sift 

through soil to check for grubs. If more 

than one grub per hole is present, a 

preplanting treatment is recommended. 

Preseason 


at this time. 


• Threshold levels are for grubs in soil prior 

to planting. See Plantation Establishment 

above. 




Biological/Biorational 

• The bacterial powder Bacillus popillae 

causes a milky spore disease specifi c to 

Japanese beetle. Application to grassy 

areas in early summer or late fall allows 

grubs to ingest the bacteria, which will 

kill them before spring. 

• Pathogenic fungi, nematodes, and protozoans 

may help decrease grub populations. 

• The parasitic wasps Tiphia popilliavora 

(Rohwer) and Tiphia vernalis (Rohwer) 

are parasites of Japanese beetle and some 

other white grubs. 

• Birds and toads consume many beetles in 

addition to a small quantity of grubs. 

• Moles, shrews, and skunks feed heavily on 

grubs in the soil. 

Mechanical 

• For low numbers of grubs, hand removal 

from tilled soil may be helpful. 

• For new seedling beds, till or disk several 

times in April–May or September to 

injure larvae in soil and expose them to 

parasite and predators. 

• Some states recommend maintenance 

of groundcover to offer alternate food 

sources to grubs in the plantation. 

Chemical 

• Apply a registered insecticide with a 

stomach poison to grassy areas in the 

spring (March–mid-May) or fall 

(September–November) to control grubs. 

• Apply a registered insecticide as a preventative 

drench at the base of the plant, 

directed at the root ball/zone. 

• Root dip may be hard on trees, so in-fi eld 

treatment is preferred. 

Figure 3. Seedling with grub 

feeding damage (left) and 

healthy seedling (right). 

Courtesy of James Solomon, 

USDA Forest Service, 


WHITE GRUBS (MAY, JUNE, AND JAPANESE BEETLES) ...................................................................................................................................................................................................................... 119

ZIMMERMAN 

PINE MOTH 

Dioryctria zimmermani 

(Grote) 

Pitch mass, a symptom of 

Zimmerman pine moth, 

located at the junction of 

a lateral branch and main 

stem. Courtesy of Whitney 

Cranshaw, Colorado State 

University, Bugwood.org 

(#1325082) 

Hosts 

• All pines, especially Austrian and Scotch 

• Occasionally found on Douglas-fi r and 

spruce 



Symptoms 

• Popcornlike, white to pinkish pitch mass 

on trunk, often at a branch whorl or on 

shoots near the terminal leader; transplants 

and young trees may have pitch 

mass closer to soil line 

• Reddish, sawdustlike frass mixed with 

pitch at entrance sites of bore holes 

• Broken leader or lateral branches; dead or 

dying branches on upper half of tree 

• Sawdust collecting on lateral branches 

and in webbing on the tree in early 

summer 

• Empty pupal case protruding from wound 

in main trunk 


• White pine weevil 


Adults, eggs, and young larvae of Zimmerman 

pine moths are very rarely seen. Their 

coloration and elusive habits help them 

avoid detection. Field identifi cation is usually 

through the presence of a pitch mass 

on the main trunk. This mass is popcornlike 

and yellow to pinkish in color. Frass is 

usually mixed in with the pitch. When the 

glob of pitch is removed, a small, elongate/ 

oval hole is usually found in the trunk. In 

an active infestation, the pitch mass will be 

somewhat pliable and soft. Hardened masses 

are generally from larval activity in the 

previous season. 

From May to August, a single larva 

may be found in a short gallery in an active 

infestation. Zimmerman pine moth larvae 

are pinkish to greenish and up to an inch 

(25 mm) long when mature. They have a 

dark brown head and numerous dark spots 


Symptoms 

Monitor 

Mechanical 

Control 

Spray Control 

on the body. The ¾-inch-long (18-mm) 

pupa may be found at the exit of the feeding 

tunnel from mid-July to late August. 


Zimmerman pine moths overwinter in the 

bark of the tree trunk as small caterpillars 

that emerged from eggs in late summer or 

early fall. The newly hatched caterpillars 

do not feed but quickly move to nearby 

protected sites under bark scales or in 

crevices below a main lateral branch. They 

chew small cavities in the bark and cover 

themselves with a silken tent, or hibernaculum, 

in which they will overwinter. 

When Scotch pine terminal growth starts 

the following April, the larvae leave their 

overwintering site and chew into the inner 

bark around the junction of a lateral branch 

and main trunk. Other favored entrance 

sites include wounds and galls caused by 

pine-pine or pine-oak gall. Small amounts 

of sawdust may be seen clinging to branches 

and in spider webs at this time. 

As the larvae feed in the phloem tissue, 

sap fl ows into the tunnel. Larvae push 

this sap—mixed with their frass and other 

debris—out of the tunnel, creating the characteristic 

pitch mass (Fig. 1). Occasionally, 

larvae will leave one feeding site and create 

a second site nearby. This feeding weakens 

the tree branches and often causes breakage 

Figure 1. Pitch 

mass made up 

of sap, frass, 

and tree debris 

that has 

been pushed 

out of the 

feeding tube 

by the larva. 

Courtesy of 

Rayanne D. 

Lehman, PDA 


STEM AND ROOT INJURY/TREE MORTALITY ... ...................................................................................................................................................................................................................................... 120

Figure 2. Larva 

feeding within 

the trunk. Courtesy 

of Rayanne D. 

Lehman, PDA 

Figure 3. Weakened branch junction due 

to Zimmerman pine moth feeding damage. 

Courtesy of Minnesota Department of Natural 

Resources Archive, Bugwood.org (#4212056) 

(Figs. 2 and 3). Feeding continues through 

the summer and the larvae are mature by 

August (Fig. 4). 

The larvae pupate inside their feeding 

tunnel, close to the entrance site, or in 

the soft pitch mass (Fig. 5). Adult moths 

emerge 2–3 weeks later, generally in late 

July through August. The short-lived adults 

are weak, nocturnal fl iers that spend most 

of their time resting on the bark of the host 

tree (Fig. 6). The coloration provides excellent 

camoufl age and they are rarely seen. In 

late summer, each mated female deposits up 

to 80 eggs on the bark of the main trunk. 

The eggs hatch in early fall. One generation 

occurs each year. 


Strategies 


• Plant resistant varieties of Scotch pine, 

especially short-needled ones from 

Greece, Turkey, and others from south 

and west Eurasia. 

• Observe proper planting practices, 

including depth of transplants. Planting 

too deep will lead to health issues. 

Preseason 

• Remove and destroy heavily damaged 

trees. Zimmerman pine moth frequently 

reinfests trees; removal of these “brood” 

trees may lessen damage to healthy trees. 


• Inspect blocks on a regular basis 

throughout the growing season. 

• Look for pitch masses on the main trunk 

or near terminal leader. In young trees, 

masses maybe closer to soil line. 

• Avoid mechanical damage to trunks that 

may increase susceptibility to this pest. 

• Chemically or mechanically control pine 

galls to decrease attractive larval feeding 

sites. 

• Growing degree days: Larvae emerge from 

overwintering sites at 121–246 GDDs. 


established. 




Biological 

• Many larval and Trichogramma egg parasitoids 

have been identifi ed in southern 

Michigan. Native parasitic wasps infested 

up to 57 percent of larvae and 45 percent 

of eggs in the study area. 

Mechanical 

• Cut out areas with pitch masses on 

the main trunk with a pocket knife or 

shearing tool and use a thin wire to 

destroy larvae in gallery. 

• Hand-prune and destroy occasional 

injured shoots. 

• Remove and destroy (by burning or 

chipping) infested trees by early July 

before adults emerge. 

Biorational 


at this time. 

Chemical 

• When larvae are abundant or repeatedly 

attacking the main trunk, apply a registered 

insecticide in early April–early May 

as the weather warms. 

— Emerging larvae are the most vulnerable 

to pesticides before boring under 

bark. 

— Use enough nozzle pressure and water 

to drench branch and trunk bark. 

Figure 4. Larva with pink-green 

body, dark brown head, and 

small, dark spots. Courtesy of 


Figure 5. Pupae close to the exit 

of the feeding tunnel. Courtesy 


Figure 6. Adult moth with 

coloration that allows it to 

blend well with the tree bark. 




ZIMMERMAN PINE MOTH .................................................................................................................................................................................................................................................................... 121

OTHER TYPES OF 

DAMAGE TO CONIFERS 

Other factors beyond disease, insect, and 

mite pests can cause signifi cant damage 

to Christmas trees that may resemble pest 

damage. These factors fall into two broad 

categories. The abiotic factors include 

those not caused by living organisms as 

well as chemical, environmental, and 

mechanical damage. The biotic factors 

are those caused by several vertebrate 

pests. Both of these types of damage are 

discussed on pages 123–129. 

OTHER TYPES OF DAMAGE TO CONIFERS .......................................................................................................................................................................................................................................... 122

Chemical Damage 

Chemical damage can result from pesticide 

applications made during the growing 

season. There are varied reasons for this type 

of damage, such as mixing incompatible 

materials, using the wrong concentration, 

applying to tender foliage, overlooking drift 

or overspray, using equipment that has previously 

contained an herbicide, or even using a 

material that is not labeled for use on Christmas 

trees (Figs. 1 and 2). The root cause of 

most chemical damage is applicator error. 

Figure 1. 

Glyphosate 

damage to spruce. 

Courtesy of 

Rayanne D. 

Lehman, PDA 

Figure 2. Distortion of spruce foliage due to 

herbicide application. Courtesy of Ricky Bates, 

Penn State 

Figure 3. Chemical burn all along the row on 

lower portion of the trees typical of improper 

sprayer adjustment. Courtesy of Rayanne D. 

Lehman, PDA 

Typically, symptoms of chemical damage 

follow the path of application through the 

fi eld (Fig. 3). If drift or overspray is thought 

to be the culprit, damage would only occur 

in the portion of the fi eld closest to the 

intended spray target (Fig. 4). If the damage 

is throughout the fi eld, mixing error, use of 

the wrong material, or equipment contamination 

may be the cause. Damage limited 

to new growth may result if an application 

is made before the growth has hardened 

off. These types of damage appear quickly 

during the growing season. During the rest 

of the year, damage appears at a slower rate. 

Some insecticide oils can burn new foliage, 

but most chemical injuries to trees are from 

herbicide applications (Figs. 5 and 6). 

Figure 4. 

Chemical 

burn due to 

improper 

sprayer 

adjustment. 

Courtesy of 

Rayanne D. 

Lehman, PDA 

Figure 5. Damage from simazine application. 

Courtesy of Ricky Bates, Penn State 

Figure 6. Trees exhibiting 

glyphosate burn on left; 

healthy trees on right. Courtesy 

of Ricky Bates, Penn State 

CHEMICAL DAMAGE ............................................................................................................................................................................................................................................................................ 123

Figure 1. Air pollution damage 

on eastern white pine. 


Service Region 8 Archive, 


Figure 2. Needle discoloration 

and drop as a result of air 

pollution. Courtesy of USDA 

Forest Service Northeastern 

Area Archive, Bugwood.org 

(#1398007) 

Environmental Damage 

Air Pollution 

Air pollution has been shown to affect all 

conifers, with each species responding differently. 

In general, eastern white pine is 

the most susceptible, but spruce may be the 

most sensitive in industrial areas. 

A tree’s response to airborne pollutants 

is affected by temperature, light intensity, 

water availability, and soil nutrients. 

Airborne pollutants may cause a decrease 

in photosynthesis, resulting in carbohydrate 

shortage in older needles. This leads to a 

decrease in new needle development and 

growth. Trees attempt to conserve energy 

when exposed to high levels of air pollution 

and may undergo early needle drop or 

senescence (Figs. 1 and 2). Trees growing in 

shallow or poorly drained soils may already 

be near the limits of their environmental 

requirements and would be predisposed to 

stress from air pollution. Trees growing in 

better soils are more susceptible to damage 

in late summer due to the high rates of 

photosynthesis at that time. Decreased 

radial growth (diameter expansion) often 

results, but tree death may also occur. 

Some experiments have shown decreases 

in tree hardiness and frost resistance due to 

an increase in ozone levels caused by pollution. 

However, others have shown that high 

ozone levels did not affect frost resistance. 

On the positive side, a few studies have 

shown that an increase in carbon dioxide 

(CO 2 ) improves photosynthesis and rootto-shoot 

ratio in fi eld-grown plants, provided 

they are of optimal nutrition. Some 

naturally occurring antioxidants have been 

identifi ed in trees. These compounds may 

reverse some of the effects of air pollution. 

From available research, it has been shown 

that conifers have a higher tolerance to 

pollution during their dormant period than 

when actively growing. 

Suggestions for limiting damage from 

air pollution include pruning dead shoots, 

culling dying trees to prevent other pests, 

avoiding planting near a source of pollution, 

and planting air-pollution-resistant species. 

Drought 

Drought damage is the result of trees needing 

more moisture than what is available 

in the soil. It affects all ages of trees, but 

new transplants are the most vulnerable. 

Symptoms generally start at the top of 

the tree and continue downward and may 

include the loss of needles, fading or yellowing 

of needles, and overall slow growth. 

During extreme drought, tree mortality may 

occur (Fig. 3). The effects of drought may 

be from one growing season of signifi cantly 

decreased moisture or from several seasons 

of below-average rainfall. Drought also leads 

to other problems, especially insect pests 

and diseases. 

Heat and Sunscald 

Extreme heat from intense sunlight can 

damage all Christmas trees, especially 

during bud break. Heat damage generally 

occurs on the southern or southwestern exposure, 

where needles quickly turn reddish 

brown, causing the entire tree to appear 

burned (Fig. 4). 

Sunscald is caused when young, thinbarked 

trees are exposed to concentrated 

sunlight and the cambium under the bark 

dies. It may also coincide with drought 

conditions. The bark will take on a coppery 

brown color during summer and branches 

above the site may die. Eventually, the bark 

may slough off. Damage may be limited to 

Figure 3. A whole block of dead seedlings as a 

result of drought. Courtesy of PDA 

Figure 4. Foliage burn on the southern side of 

the tree caused by intense sunlight. Courtesy of 



a certain area or may occur over the entire 

tree but is generally restricted to the southern 

or southwestern exposure. Trees with 

few branches are more likely to experience 

sunscald. Douglas-fi r is the most susceptible 

to sunscald, with true fi rs following closely 

behind (Fig. 5). Basal pruning in fall or 

winter and commercial tree wraps may help 

reduce this problem. 

Frost Injury 

Frost injury occurs in early spring and is most 

common on Douglas-fi r, true fi rs, and spruce. 

Pines are damaged only on rare occasions 

of extremely late season frost. Trees that 

break bud early or are in frost-prone areas 

on northern slopes are most susceptible to 

this type of damage. Within a few days of the 

frost event, the new growth wilts and turns 

pinkish brown. As the season progresses, 

damaged shoots turn dark brown and may 

remain on the tree into summer (Fig. 6). 

Damaged growth can be removed during 

shearing, and the tree should recover within 

one year. Repeated frost damage will result 

in trees that are underdeveloped and bushy. 

Pines are best suited for planting in known 

frost pockets or on northern exposures. 

Figure 5. Douglas-fi r showing symptoms of 

sunscald. Courtesy of USDA Forest Service 

Ogden Archive, Bugwood.org (#1467233) 

Figure 6. New 

growth of Fraser 

fi r damaged by 

frost. Courtesy of 

Ricky Bates, Penn 

State 

Winter Injury 

Winter injury is more common with Fraser 

and Concolor fi r, eastern white pine, and 

Norway spruce. Both winter burn and drying 

may occur, causing the needles to turn 

brown. Occasionally, an entire tree will 

be discolored above the snow line (Figs. 7 

and 8). Needle loss may occur, especially 

on the southern side. On eastern white 

pine, winter injury causes needles to droop 

downward. On Scotch pine, yellow spots are 

found on damaged needles. 

Wind Damage 

Wind damage often occurs on the northwest 

or west side of the trees. It is also more 

common around the perimeter of the tree 

block. Needles will show discoloration from 

the tips of the needles to the back (Fig. 9). 

On closer inspection, needles may have a 

stippled effect. 

Figure 7. Snow 

protecting foliage 

from extreme 

winter elements. 

Courtesy of 

PDA 

Figure 8. 

Foliage that 

sat above 

the snow 

line exhibiting 

dieback. 

Courtesy of 

Rayanne D. 

Lehman, PDA 

Delayed Winter Dormancy 

Delayed winter dormancy may occur when 

climate conditions prior to and at the time 

of harvesting Christmas trees are warmer 

and/or wetter than usual. Typically, growers 

in the East begin harvesting trees in mid- to 

late November. When November temperatures 

are warmer than usual, trees may not 

have the opportunity to become dormant 

before being cut. When this occurs, trees are 

more likely to dry out and prematurely drop 

needles. Delaying tree cutting until after 

temperatures have dropped may be a way to 

prevent early needle loss. 

Figure 9. Tree near the perimeter 

of a block exhibiting wind 

damage. Courtesy of Rayanne D. 

Lehman, PDA 

ENVIRONMENTAL DAMAGE ................................................................................................................................................................................................................................................................ 125

Figure 1. Exhaust burn at 

the bottom of a tree. 


Mechanical Damage 

Growing Christmas trees requires the 

use of many different types of equipment 

and tools. If not properly used, these may 

actually damage the trees. 

Equipment Damage 

Equipment used in a new tree block generally 

passes trouble free down the straight 

rows. As the trees grow, causing the rows to 

narrow, it becomes harder for the equipment 

to travel down the rows without causing 

damage. Equipment damage results from the 

tractor and its tires and implements. 

Equipment used for mowing and spraying 

can also cause damage to the trees. 

Entire small trees can be crushed and lower 

branches of larger trees can be broken or 

pulled from the trunk by equipment tires. 

The spray boom can scrape branches as 

well as damage bark and buds. Mowers are 

known to scrape the base of trees. Trees at 

the ends of the rows are more susceptible to 

damage because of turning-radius requirements 

by the tractor and implements. It 

may take several weeks after the damage 

occurs for symptoms to appear, but they are 

most quickly noticeable around bud break. 

Equipment damage is often permanent and 

disfi guring. 

Some growers have modifi ed their equipment 

by adding guards to prevent branches 

of trees from being pulled into the path of 

the tires or implements. Damage can also be 

reduced by widening the distance between 

rows. Where the current standard is 5 feet 

by 5 feet (1.52 by 1.52 m), some farms are 

going to 5 feet by 6 feet (1.52 by 1.83 m) or 

even 6 feet by 6 feet (1.83 by 1.83 m). It is 

important to provide ample turning area at 

the end of rows when planting. 

Heat and fumes from equipment exhaust 

can damage trees. Exposure time does not 

have to be long, especially with new growth. 

Symptoms include discolored or browned 

needles and dying shoots (Fig. 1). The 

damaged area is generally about one square 

foot. Exhaust damage can be diagnosed by 

looking down the rows. If exhaust damage is 

likely, the burned spots will be at the same 

height on numerous trees in the row. Narrow 

rows are most susceptible. 

A few solutions for reducing this type of 

damage include directing exhaust away from 

trees, keeping hot engine parts away from 

trees, using a catalytic converter, and keeping 

equipment speeds increased rather than 

decreased. 

String trimmers cause serious damage to 

trees by damaging the bark at the base of the 

tree. This is especially true when trees are 

larger and the operator has diffi culty seeing 

into the trunk. The cuts and gouges in the 

bark offer entrance sites for several insect 

pests and diseases. To prevent string trimmer 

damage, maintain good weed control in the 

rows and avoid use of string trimmers under 

trees. 

Planting equipment may also lead to tree 

injury. If the planting depth is not correct, 

the taproot may curve upward instead of 

straight down. The tree roots will continue 

to grow upward—a condition known as 

J-rooting—and the trees will become 

stunted and eventually die (Fig. 2). To avoid 

J-rooting, adjust mechanical planters for 

each use with the proper planting depth required 

for the size of transplants being used. 

Figure 2. A J-rooted tree. Courtesy of PDA 


Shearing Damage 

Shearing can damage trees. If a shearing 

knife is not used properly or is not sharp, 

branches will not be cut cleanly and partial 

cuts or broken branches may result. This 

results in dead or “fl agged” branches. To 

reduce damage when using a shearing knife, 

always use a sharp blade and swing with 

enough force to ensure minimal uncut 

branch ends (Fig. 3). 

Improper machine shearing can also 

harm trees. If the branches move away from 

the moving teeth, they will be left with a 

“chewed off” look. The end result is yellow, 

unattractive, or dead branch ends. Trees 

can be damaged by excess lubricating oil 

and will appear burned. Avoid using too 

much when lubricating the blades. When 

machine shearing, keeping the equipment 

at the proper angle and speed while cutting 

into the branches will reduce the “chewed 

off” look of branch ends. 

Shearing is dangerous, so use all recommended 

safety equipment and provide 

training to those working on this activity. 

Figure 3. Clean, sharpened, 16-inch shearing 

knife. Courtesy of Sarah Pickel, PDA 

MECHANICAL DAMAGE ....................................................................................................................................................................................................................................................................... 127

Figure 1. Bird perching on a 

tree, which can cause the 

leaders to bend or break. 

Courtesy of Rayanne D. 

Lehman, PDA 

Vertebrate Pests 

Birds 

Birds tend not to be a big problem associated 

with Christmas tree production, but damage 

occasionally occurs to the tops, particularly 

of taller trees (Fig. 1). This is especially common 

during spring, when the new growth 

is supple and not hardened off. The weight 

of the birds causes the terminal to bend and 

break. This damage is sometimes mistaken 

for white pine weevil damage; however, the 

birds will cause a sharp bend or break instead 

of the curled, wilted damage caused by the 

weevil. Taller trees are targeted because birds 

need a good view in order to defend their 

territory. Most growers having repeated damage 

will install poles or perches above tree 

height to alleviate the problem. 

Later in the year, birds may cause 

another type of damage as they perch on 

the stiff, mature terminals. Needles may be 

pulled from the terminal by the birds’ feet. 

This damage is minimal and will be corrected 

with another year’s growth. Noiseproducing 

devices have been used, but birds 

learn to tolerate them over time. 

Deer 

Deer will feed on all conifers, but spruce 

is the least preferred. Feeding damage may 

occur during winter when preferred foods 

are not available. The ends of deer-browsed 

shoots appear ragged because deer lack upper 

incisors and must bite down and rip food 

sources in order to feed (Fig. 2). 

Figure 2. Damage from deer browsing. 

Courtesy of John H. Ghent, USDA Forest 


The most common damage from deer 

occurs in late summer and fall when bucks 

polish their antlers by rubbing them against 

the trunks of small trees (Fig. 3). Damage 

may be more evident along the edge of 

blocks adjacent to woodlots as deer are an 

edge species. White-tailed deer cause more 

damage to Christmas trees than any other 

animal. 

Methods of limiting deer damage to 

Christmas trees include fencing and repellents. 

Fencing must be justifi able by being 

economical and of low maintenance. Studies 

on types of deer fence have shown three 

kinds to be effective. Woven wire fence is 

the best choice where deer damage is intolerable. 

Multistrand, electric, high-tensile 

fences are good in situations when total 

deer exclusion is not a necessity and the 

cost of fencing is not an issue. An electric 

poly-fence is good for seasonal protection. 

Each grower must look at all fencing options, 

farm operations, fi nances, and goals to 

decide the best choice for their farm. 

A second possible option for limiting 

deer damage is the use of repellents. Feed 

treated with lime provided the best repellency 

in one study; charcoal was the second 

most effective. Bittering agents were shown 

not to be effective over extended periods in 

another study. Growers have hung bars of 

Ivory soap from the trees with mixed results. 

Some other options for deterring deer 

are keeping a dog in the vicinity of the tree 

fi elds and allowing hunting on or near the 

property. 

Figure 3. Conifer bark and branches 

damaged by bucks polishing their antlers. 

Courtesy of Sarah Pickel, PDA 


Mice and Voles 

Damage to trees from these rodents is highest 

in the winter and early spring when a 

shortage of their preferred foods is present. 

During winter, both mice and voles will feed 

under the snow cover and consume bark 

cambium and phloem layers and the roots 

of trees (Fig. 4). A decrease in tree growth 

results from the sublethal feeding injuries, 

but smaller trees may die if completely 

girdled (Fig. 5). 

The best way to control these rodents 

is by maintaining weed control in the rows 

and particularly around the base of the tree. 

The bare ground is not attractive to the 

rodents and exposes them to predators. 

Figure 4. Rodent feeding on root tissue. 


Figure 5. Girdling at the base of the tree 

from rodent feeding. Courtesy of Rayanne D. 

Lehman, PDA 

Rabbits 

Rabbits will feed on young trees. Shoots 

cut off at a 45-degree angle or girdling at 

the base are symptoms of rabbit feeding. 

Damage frequently occurs in winter when 

rabbits are unable to reach their normal 

food sources because of snow cover. During 

these periods, they will feed higher on the 

tree and may remove signifi cant amounts 

of bark. Good weed control in the rows and 

around the trees removes hiding places for 

rabbits and makes them more vulnerable to 

predation. 

Groundhogs 

Groundhogs can do minimal chewing damage 

to tree bark, but the bigger problem is 

their burrows. These holes are hazardous 

to farm equipment and employees on foot. 

When groundhogs burrow under a tree, 

the root system is disturbed and the tree 

may be weakened. Maintaining good weed 

control, either chemically or by mowing, 

and eliminating brush piles will discourage 

groundhogs in the fi eld. Some recommendations 

include live trapping to relocate, 

smoke bombs, coyote or fox urine around 

the burrow, or motion devices to frighten 

these timid but destructive animals. 

VERTEBRATE PESTS .............................................................................................................................................................................................................................................................................. 129

PESTICIDE INFORMATION 

On most Christmas tree farms, pesticides play 

a role in management of pest issues. Pesticides 

are chemicals used to kill, mitigate, or repel 

pests; as such, both benefi ts and dangers may 

be associated with their use. To prevent misuse 

and undesirable effects, the government has 

enacted regulations on the use and handling of 

pesticides. This section discusses some of those 

pesticide regulations. 

Information on the safe use of pesticides 

can be found through county cooperative 

extension offi ces, state land-grant universities, 

state agricultural departments, or the United 

States Environmental Protection Agency. In 

Pennsylvania, Penn State’s Pesticide Education 

Program provides a wealth of educational 

materials. This information can be found at 

www.pested.psu.edu. 

PESTICIDE INFORMATION .................................................................................................................................................................................................................................................................... 130

Fungicides, Herbicides, 

Insecticides, and Miticides 

Pesticides, by defi nition, are chemicals or 

other agents used to kill or control pests or 

to protect something from a pest. Generally, 

four types of pesticides are used in the Christmas 

tree industry: fungicides used to control 

disease-causing fungi; herbicides to kill or 

deter plant growth; insecticides to control 

or prevent damage caused by insects; and 

miticides, often associated with insecticides, 

to control mites or arachnids (spiders and 

ticks). Each pesticide can be identifi ed by its 

brand name, active ingredient, and chemical 

class. Closely related chemicals are grouped 

in the same chemical class. 

Pesticide Resistance Issues 

One major concern that has been raised as 

the agricultural industry becomes increasingly 

dependent on pesticides is the risk of 

pests, including diseases and weeds, developing 

resistance to current pesticides. Repeated 

use of the same class of pesticide hastens this 

development and, ultimately, renders the 

chemical ineffective. This is especially true 

for pests that have multiple generations each 

growing season. By regularly changing to a 

compound from a different chemical class, 

the development of resistance is slowed, thus 

extending the useful life of the pesticide. 

Several international groups have 

formed to address the issue of pesticide 

resistance: 

• The Fungicide Resistance Action Committee 

(FRAC) has developed a number 

and letter code to distinguish the fungicide 

groups according to their cross-resistance 

behavior. A table of codes can be found at 

www.frac.info/frac/publication/anhang/ 

FRAC_CODE_LIST.pdf. 

• The Herbicide Resistance Action 

Committee (HRAC) supports the global 

survey of resistant weeds initiated by the 

Weed Science Society of America. A 

searchable database of resistant weeds 

can be found at www.weedscience.com. 

• The Insecticide Resistance Action Committee 

(IRAC) has developed a mode 

of action (MOA) classifi cation based on 

known ways in which different products 

act. The MOA Classifi cation list can be 

downloaded from www.irac-online.org/ 

News.asp. 

Pesticide Safety 

Federal and Pennsylvania Pesticide Laws 

The use of pesticides worldwide has increased 

greatly over the last 35 years. 

With this increased use, there are growing 

concerns for possible harm to human 

health and the environment. The purpose 

behind pesticide regulation is to protect 

public health and welfare, as well as prevent 

harmful effects on the environment through 

proper labeling, sale and distribution, transportation, 

storage, use and application, and 

disposal of pesticides. Pesticides are under 

regulatory inspection from the time they 

are invented in the lab to use in the fi eld or 

approved disposal. 

Since signed into law in 1947, the Federal 

Insecticide, Fungicide, and Rodenticide Act 

(FIFRA) has been the basis for regulation, 

distribution, and use of pesticides in the 

United States. FIFRA grants the federal 

Environmental Protection Agency the power 

to stop the sale or use of any pesticide. FIFRA 

regulations dictate labeling and packaging 

standards and also disposal procedures for 

pesticide containers and surplus pesticides. 

The Pennsylvania Pesticide Control Act 

was enacted in 1973 as a companion bill to 

FIFRA. The purpose of this state act is to 

regulate the labeling, distribution, storage, 

transportation, use, application, and 

disposal of pesticides. In addition to being 

registered with EPA, every pesticide used in 

Pennsylvania must also be registered with 

the Pennsylvania Department of Agriculture. 

Regulations also cover dealers and 

applicators and the licensing of applicators. 

For additional information, contact the 


at www.paplants.state.pa.us/Index.aspx or 

Penn State’s Pesticide Education Program at 

www.pested.psu.edu. If you reside outside 

Pennsylvania, consult your local regulatory 

authority or land-grant university. 

Worker Protection Standard 

In August 1992, the Worker Protection 

Standard (WPS) was revised. Like FIFRA, 

it is overseen by the U.S. Environmental 

Protection Agency. This regulation is aimed 

at decreasing the risk of pesticide poisoning 

and injuries to agricultural workers and 

pesticide handlers. Included in the WPS are 

requirements for pesticide safety training, 

worker notifi cation of pesticide applications 

in the workplace, personal protective equipment 

use, reentry interval, decontamination 

supplies, and emergency medical assistance. 

CAUTION: 

Always read the 

pesticide label to 

determine specifi c 

uses and rates before 

mixing and applying 

the compound. If 

any questions arise, 

contact the dealer or 

manufacturer. It is 

illegal to apply any 

pesticide in excess of 

labeled rates. Labeled 

uses may vary for 

each formulation of 

the same chemical. 

Purchase the 

formulation intended 

for your particular use. 

PESTICIDE INFORMATION .................................................................................................................................................................................................................................................................... 131

APPENDIXES

APPENDIX A 

PEST AND DISEASE 

PHOTO CHART 

This photographic appendix is designed to 

ease identifi cation of disease and insect pest 

problems. Two photographs are provided for 

each pest, which are grouped according to 

the type of damage they represent. The fi rst 

photograph is an overall view of the damage 

and/or symptoms, while the second photograph 

is a close-up of the pest problem or 

the actual pest causing the symptoms. The 

fi rst column, “Name of Pest,” states what 

the problem is. The second column, “Trees 

Susceptible,” indicates which trees are hosts 

for the particular pest. The page number of 

the appropriate pest fact sheet is provided 

in the third column. 

APPENDIX A ...................................................................................................................................................................................................................................................................................................................... APPENDIX A .......................................................................................................................................................................................................................................................................................... 134 134

Pest and Disease Photo Chart 

Name of Pest Trees Susceptible Page Name of Pest Trees Susceptible Page 

Bagworm Douglas-fi r, fi r, pine, spruce 23 Cyclaneusma needle cast Pine 34 

S. GARDOSIK, PDA 

Balsam twig aphid Fir 26 

T. OLSON, PDA 

Cooley spruce gall adelgid Douglas-fi r 29 

(on Douglas-fi r) 


Cryptomeria scale Douglas-fi r, fi r, pine, spruce 31 


C. THOMAS, PDA 

R. LEHMAN, PDA 



NEEDLE DISCOLORATION AND INJURY 

USDA FOREST SERVICE 

Douglas-fi r needle midge Douglas-fi r 37 

Elongate hemlock scale Douglas-fi r, fi r, pine, spruce 39 

PEST AND DISEASE PHOTO CHART ................................................................................................................................................................................................................................................................................. 135 

T. OLSON, PDA 

S. GARDOSIK, PDA S. GARDOSIK, PDA 

T. OLSON, PDA 

S. GARDOSIK, PDA


Eriophyid rust and sheath mites Pine, fi r, spruce 42 Introduced pine sawfl y Pine 53 


European pine sawfl y Pine 53 


Gypsy moth Douglas-fi r, fi r, pine, spruce 45 

PDA 


NEEDLE DISCOLORATION AND INJURY (continued) 

S. KATOVICH, USDA FOREST SERVICE 

B. SCHILDT, PDA 

J. H. GHENT, USDA FOREST SERVICE 

Lophodermium needle cast Pine 48 

E. L. BARNARD, FLORIDA DEPARTMENT OF AGRICULTURE AND 

CONSUMER SERVICES 

Pine needle scale Douglas-fi r, fi r, pine, spruce 51 



J. H. GHENT, USDA FOREST SERVICE 

Ploioderma needle cast Pine 56 

APPENDIX A ...................................................................................................................................................................................................................................................................................................................... 136 

T. OLSON, PDA 

T. OLSON, PDA 

T. OLSON, PDA


Red-band (Dothistroma) 

needle blight Pine 58 

T. OLSON, PDA 

Redheaded pine sawfl y Pine 53 


T. OLSON, PDA 

Rhabdocline needle cast Douglas-fi r 60 

T. OLSON, PDA T. OLSON, PDA 

Rhizosphaera needle cast Spruce 63 

T. OLSON, PDA 



Spruce needle rust Spruce 65 

Spruce spider mite Douglas-fi r, fi r, pine, spruce 67 

Swiss needle cast Douglas-fi r 70 


T. OLSON, PDA 

E. R. DAY, VIRGINIA TECH 

T. OLSON, PDA 

T. OLSON, PDA 


T. OLSON, PDA


Atropellis canker Pine 73 Cooley spruce gall adelgid 

(on Colorado blue spruce) Spruce 80 

T. OLSON, PDA 

Balsam woolly adelgid Fir 75 


Botrytis blight Douglas-fi r, fi r, pine, spruce 78 

T. OLSON, PDA 

T. OLSON, PDA 


T. OLSON, PDA 

SHOOT AND BRANCH INJURY 



Diplodia (Sphaeropsis) tip blight Douglas-fi r, pine, spruce 82 

Eastern pine weevil Douglas-fi r, pine, spruce 84 


APPENDIX A ...................................................................................................................................................................................................................................................................................................................... 138 

T. OLSON, PDA 

W. JOHNSON, USDA FOREST SERVICE 

T. OLSON, PDA


Eastern spruce gall adelgid Spruce 86 Pine shoot beetle Pine 94 

S. GARDOSIK, PDA R. LEHMAN, PDA 

Gall rusts Pine 88 

T. OLSON, PDA T. OLSON, PDA 

Pales weevil Douglas-fi r, pine, spruce 90 


Pine bark adelgid Pine 92 





Striped pine scale Pine 97 




White pine blister rust Pine 99 

White pine weevil Douglas-fi r, fi r, pine, spruce 101 


T. OLSON, PDA 

PDA 

T. OLSON, PDA 

S. KATOVICH, USDA FOREST SERVICE


Armillaria root rot Douglas-fi r, fi r, pine, spruce 107 Phytophthora root rot Fir, pine, spruce 111 

J. O’BRIEN, USDA FOREST SERVICE 

Bark and engraver beetles Pine 109 

PDA 

J. STIMMEL, PDA 

Japanese beetle Douglas-fi r, pine, spruce 118 

STEM AND ROOT INJURY/TREE MORTALITY 

D. POWELL, USDA FOREST SERVICE 



Pine root collar weevil Pine 113 

Pine wilt disease Pine 116 

APPENDIX A ...................................................................................................................................................................................................................................................................................................................... 140 

T. OLSON, PDA 


A. S. MUNSON, USDA FOREST SERVICE 

T. OLSON, PDA 

J. B. HANSON, USDA FOREST SERVICE 

L. D. DWINELL, USDA FOREST SERVICE


White grubs Douglas-fi r, fi r, pine, spruce 118 Zimmerman pine moth Pine, spruce 120 


Air pollution 

(environmental damage) Douglas-fi r, fi r, pine, spruce 124 

MINNESOTA DEPARTMENT OF NATURAL RESOURCES 

S. KATOVICH, USDA FOREST SERVICE 

J. STIMMEL, PDA 

OTHER TYPES OF DAMAGE TO CONIFERS 

MINNESOTA DEPARTMENT OF NATURAL RESOURCES 

W. CRANSHAW, COLORADO STATE UNIVERSITY 


Drought 


Chemical damage Douglas-fi r, fi r, pine, spruce 123 Frost injury 


R. BATES, PENN STATE 

T. OLSON, PDA 

W. M. BROWN JR. 


R. L. ANDERSON, USDA FOREST SERVICE 


PEST AND DISEASE PHOTO CHART ................................................................................................................................................................................................................................................................................. 141


Heat (environmental damage) Douglas-fi r, fi r, pine, spruce 124 Vertebrate pests Douglas-fi r, fi r, pine, spruce 128 


Mechanical damage Douglas-fi r, fi r, pine, spruce 126 

PDA 

Sunscald 

(environmental damage) Douglas-fi r, fi r 124 

USDA FOREST SERVICE OGDEN 

PDA 


S. K. HAGLE, USDA FOREST SERVICE 

OTHER TYPES OF DAMAGE TO CONIFERS (continued) 

USDA FOREST SERVICE NORTH CENTRAL RESEARCH STATION 

Winter injury 




E. G. VALLERY, USDA FOREST SERVICE 

APPENDIX A ...................................................................................................................................................................................................................................................................................................................... 142

APPENDIX B 

BIOLOGICAL CONTROLS 

PHOTO CHART 

This photographic appendix is designed 

to help with identifying biological 

controls, or biocontrols (both insects 

and fungi). The fi rst column, “Targeted 

Pest(s),” indicates pest(s) on which 

each biocontrol will prey. The common 

name and a photograph of each biocontrol 

appear in the second column, 

which is arranged alphabetically by 

biocontrol common name. The third 

column provides the scientifi c name 

of the biocontrol being viewed. 

APPENDIX B APPENDIX ...................................................................................................................................................................................................................................................................................................................... B .......................................................................................................................................................................................................................................................................................... 144 144

Targeted Pest(s) Biological Control Scientifi c Name 

Cryptomeria scale Aphytis (parasitoid) wasp Aphytis spp. 

Aphids (balsam 

twig, etc.); adelgids 

(balsam woolly, pine 

bark, Cooley spruce 

gall); other small, 

soft-bodied insects 

Wood borers, woodinhabiting 

beetles 

S. PICKEL, PDA 

Biological Controls Photo Chart 

Brown lacewing Hemerobius stigma 

Stephens 


Checkered beetle Cleridae family 


Scales Cn beetle (predatory beetle) Cybocephalus 

nipponicus 

Endrödy-Younga 



Aphids, beetle larvae, 

caterpillars (small), 

moths, spider mites 

Douglas-fi r needle 

midge 

Common green lacewing Chrysoperla 

rufi labris 

(Burmeister) 

Crab spider Thomisidae family 

Mite eggs and adults Dusty wing Coniopterygidae 

family 

Adelgids, aphids, 

adult beetles, beetle 

larvae, caterpillars, 

fl ies, mites, other pest 

insects 



C. MOOREHEAD 

Entomopathogenic fungi Isaria sp. 

M. BARBERCHECK, PENN STATE 

Aphids Eye-spotted lady beetle Anatis mali (Say) 


BIOLOGICAL CONTROLS PHOTO CHART ......................................................................................................................................................................................................................................................................... 145


Aphids Fifteen-spotted lady beetle Anatis labiculata 

(Say) 

Aphids Golden-eyed lacewing Chrysopa oculata 

(Say) 

Aphids Hudsonian lady beetle Mulsantina 

hudsonica (Casey) 

Aphids, insect eggs, 

misc. larvae/nymphs, 

mites 

A. BATEMAN, BUGGUIDE.NET 

CLEMSON UNIVERSITY, USDA COOPERATIVE EXTENSION 

SLIDE SERIES 

J. BAILEY, BUGGUIDE.NET 

Insidious fl ower bug, minute pirate 

bug 


Orius spp. 

Armored scales Lady beetle Microweisia spp. 

R. COWLES, CONNECTICUT AGRICULTURAL EXPERIMENT 

STATION 

Scales Lady beetle (white ladybug larva) Hyperaspis spp. 



Adelgids (exposed 

stages), aphids, mites 

Lady beetle Stethorus spp. 

Scales Lindorus beetle Lindorus 

lophanthae 

(Blaisd.) 

Various insects Mantids Mantidae family 

Aphids, scale insects, 

other soft-bodied 

insects 

F. PEAIRS, COLORADO STATE UNIVERSITY 

RINCON-VITOVA INSECTARIES 


Multicolored Asian lady beetle Harmonia axyridis 

(Pallas) 

PENNSYLVANIA DCNR 

Armored scales Parasitic wasp Encarsia citrina 

(Crawford) 

R. COWLES, CONNECTICUT AGRICULTURAL EXPERIMENT 

STATION 

Aphids Pine lady beetle Mulsantina picta 

(Randall) 

C. MOOREHEAD 

APPENDIX B ...................................................................................................................................................................................................................................................................................................................... 146


Caterpillars, sawfl y 

larvae 

Potter wasp Eumenes fraternus 

Say 

Spider mites Predatory mite Neoseiulus fallacis 

(Garman) 

Hemlock woolly 

adelgid 


larvae, other insects 


larvae 

R. BROEKHUIS, BUGGUIDE.NET 

NORTH CAROLINA STATE UNIVERSITY 

Pt beetle (lady beetle) Pseudoscymnus 

tsugae Sasajii and 

McClure 

D. J. SOUTO, USDA FOREST SERVICE 

Shield/stink bug Pentatomidae 

family 

R. L. ANDERSON, USDA FOREST SERVICE 

Sphecid wasp Ammophila spp. 

J. LAWRENCE, EUROFINS AGROSCIENCE SERVICES 


Aphids, other softbodied 

insects 

Syrphid fl y (hover fl y) Syrphidae family 

Aphids Three-banded lady beetle Coccinella 

trifasciata 

Linnaeus 

Aphids Transverse lady beetle Coccinella 

transversoguttata 

Falderman 

Aphids, scales Twice-stabbed lady beetle Chilocorus orbus 

Casey 

Immature and adult 

insects 


T. MURRAY, BUGGUIDE.NET 



Yellow jacket/wasp Vespula spp. 


BIOLOGICAL CONTROLS PHOTO CHART ......................................................................................................................................................................................................................................................................... 147

APPENDIX C 

CONIFER SPECIES 

These conifer species are mentioned as 

pest hosts in this manual or are common 

tree species grown in the Northeast. 

Conifers sold as Christmas trees are 

highlighted. 

APPENDIX C .......................................................................................................................................................................................................................................................................................... 148

Conifer Species 

Genus (Group) Common Name Species Name 

Abies (true fi rs) Balsam fi r Abies balsamea (L.) Mill 

Canaan fi r Abies balsamea var. phanerolepis (Fern.) 

Concolor fi r or white fi r Abies concolor (Gord. and Glend.) Lindl. 

Fraser fi r Abies fraseri (Pursh) Poir. 

Grand fi r Abies grandis (Douglas ex D. Don) Lindl. 

Korean fi r Abies koreana (Sieb and Zucc.) 

Momi fi r Abies fi rma (Sieb and Zucc.) 

Noble fi r Abies procera Rehder 

Nordmann fi r Abies nordmanniana (Steven) Spach 

Turkish fi r Abies bournmuelleriana (Mattf.) 

Juniperus (junipers) Eastern red cedar Juniperus virginiana L. 

Picea (spruce) Black hills spruce Picea glauca var. densata Bailey 

Colorado blue spruce Picea pungens Engelm. 

Englemann spruce Picea engelmannii Parry ex Engelm. 

Norway spruce Picea abies (L.) H. Karst. 

Serbian spruce Picea omorika (Pancic) Purk. 

Sitka spruce Picea sitchensis (Bong.) Carrière 

White spruce Picea glauca (Moench) Voss 

Pinus (pines) Austrian pine Pinus nigra J. F. Arnold 

Eastern white pine Pinus strobus L. 

Jack pine Pinus banksiana Lamb. 

Lodgepole pine Pinus contorta Douglas ex Louden 

Monterey pine Pinus radiata D. Don 

Mugo pine Pinus mugo Turra 

Ponderosa pine Pinus ponderosa P. Lawson and C. Lawson 

Red pine Pinus resinosa Aiton 

Scotch pine Pinus sylvestris L. 

Virginia pine Pinus virginiana (Mill.) 

Western white pine Pinus monticola (Douglas ex D. Don) 

Pseudotsuga (Douglas-fi r) Douglas-fi r Pseudotsuga menziesii (Mirb.) Franco 

Thuja (cedar) Arborvitae or northern white cedar Thuja occidentalis L. 

Tsuga (hemlock) Eastern hemlock Tsuga canadensis (L.) Carrière 

Conifers sold as Christmas trees are highlighted. 

CONIFER SPECIES .................................................................................................................................................................................................................................................................................. 149

APPENDIX D 

SEASONAL MONITORING GUIDE 

A good scouting program for your Christmas tree 

farm can lead to a more accurate pest and disease 

control program. Set time aside to scout for new 

insect and disease problems several times a year. 

Also take time to intensively scout for known 

pests to best determine the need for and timing 

of controls. As a rule, every time you are in 

a block of trees for planting, spraying, shearing, 

mowing, or harvesting, make note of any insect 

or disease issues. Combine these scouting tips 

with the control recommendations in the corresponding 

pest fact sheets found in this manual. 

Be prepared with the proper scouting tools 

before beginning these scouting activities. The 

fi rst and most important tool is a hand lens of 

15X magnifi cation or higher. Other tools include 

pruners to clip branches for sampling, plastic 

bags or vials to collect samples, and a pencil or 

pen and paper to record your fi ndings. 

Disclaimer: The suggested timing of the following 

scouting recommendations are based on 

general pest activity times in the Mid-Atlantic 

region. The best way to adapt this calendar to 

your region of the country is to follow the growing 

degree day timings listed in the fact sheet 

section of this manual. Also, the specifi c tree 

varieties on your farm may require that your 

scouting activities include pests not listed in 

this section. 

APPENDIX D .......................................................................................................................................................................................................................................................................................... 150

Seasonal Monitoring Guide 

Time of Year Hosts Pest How to Scout 

Late winter to early 

spring (March to 

early April), before 

bud break 

True fi rs Balsam twig aphid • Look for trees with curled last season’s needles. 

• On undamaged shoot, look for silvery, 

oval-shaped eggs on underside of twig within 

1–2 inches of buds. 

• Light green stem mothers feed on undersides of 

needles, excreting honeydew droplets. 

Colorado blue spruce Cooley spruce gall adelgid • Look for brown, pinecone-like galls from the 

previous season. 

• Gray-black overwintering nymphs are found at 

the base of the new season’s buds. 

Douglas-fi r Cooley spruce gall adelgid • Look for crooked needles. 

• Black nymphs will be on undersides of needles 

(may be covering over with white wax). 

Norway spruce Eastern spruce gall adelgid • Look for brown galls at the base of previous 

year’s growth; needles may be missing beyond 

gall. 

• The black overwintering nymphs will be found 

at the base of the new season’s buds. 

Pines, spruces, true fi rs Eriophyid mites • Look at 20 trees per acre that exhibit a gray or 

rusty coloring. 

• Tiny, peach-colored eggs are found in clusters or 

rows at the needle base (underside). 

• Pale, oblong adult mites move along the 

needles. 

Pines Pales weevil • Look for trees showing “fl agging,” or browning, 

of needles at the ends of branches that 

have shown signs of chewing on stem bark. 

• Pull duff away from base of last year’s stumps to 

look for adult weevils that are becoming active. 

Douglas-fi r Rhabdocline needle cast • Scout trees in areas likely to hold extra moisture 

such as shaded areas along a woodlot or tight 

plantings. 

• Examine current season’s needles for orangerust 

discoloration. 

Spruces Rhizosphaera needle cast • On the lower half of the tree, look for purplebrown 

needles with visible black fruiting bodies. 

Spruces Spruce needle rust • On cloudy days, look for needles with yellow 

bands containing an orange spot. 

• Focus scouting in low or shaded areas. 

Arborvitae, Douglas-fi r, 

spruces, true fi rs 

Spruce spider mites • Look for discolored (brown or yellow) needles. 

• Red eggs are found on the undersides of twigs, 

needles, and at base of buds. 

• Reddish-brown adult mites move along needles 

and twigs. 

• Tap symptomatic branches over a white surface 

to dislodge eggs or mites. 

Douglas-fi r Swiss needle cast • In shady or tightly planted areas, look for trees 

with needles that are browning from the tip 

down. 

• On the underside of these needles, rows of tiny, 

black fruiting bodies will be visible with a hand 

lens. 

• Also look for fruiting bodies on healthy, 

green needles. 

(continued) 

SEASONAL MONITORING GUIDE ........................................................................................................................................................................................................................................................ 151

Seasonal Monitoring Guide (continued) 

Time of Year Hosts Pest How to Scout 

Late winter to early 

spring (March to 

early April), before 

bud break 

(continued) 

Midspring to early 

summer (early April 

to June) 

Harvest (November 

to December) 

Douglas-fi r, pines, 

spruces 

Arborvitae, Douglas-fi r, 

pines, spruces, true fi rs 

White pine weevil • Set out baited pyramidal traps (see Appendix E 

for instructions). 

• Check soil temperature daily on unshaded side 

of a tree in susceptible block. 

• Once soil temperatures near 50°F, check traps 

daily. 

• On sunny days, look for sap droplets on tree 

leaders as feeding evidence. 

Bagworm • Look for trees with brown, conelike bags, 

1–2 inches in length hanging from the previous 

season’s growth. 

• Near the end of May, begin to look for tiny 

larvae feeding on the new season’s needles. 

True fi rs Balsam woolly adelgid • Look at the tops of trees for crooked leaders. 

• On symptomatic trees, try to bend the main 

trunk. Trees with balsam woolly adelgid will be 

extra stiff. 

• Branches will have gouty enlargements, or 

“warts.” 

• Small, purplish-black insects will be under the 

white, woolly wax on the bark. 

Douglas-fi r, spruces, true 

fi rs 

Cryptomeria scale • Lift the lowest branches of select trees to look 

for yellow speckling on top of needles. 

• Examine underside of needles for off-white, 

oval-shaped scales with yellow centers. Look 

closely with hand lens for moving, yellow, 

oblong crawlers among the scales. 

• In August, monitor for the second generation. 

Douglas-fi r Douglas-fi r needle midge • Prior to bud break, set out cardboard emergence 

traps (see Douglas-fi r needle midge 

fact sheet) at the base of trees with kinked or 

damaged needles from the previous season. 

• As buds swell, monitor traps for delicate, orange 

midges in the capture jar. 

• Emerging midges will also be found on breaking 

buds or hovering close by. 

Douglas-fi r, hemlocks, 




Elongate hemlock scale • Look for yellowing needles with a gray cast at 

the base of the trees, near the trunk. 

• Examine the underside of the needles for 

mobile, yellow, oblong crawlers among the 

narrow, brown scales and shorter, white scales. 

• Tapping the branch over a white paper or 

clipboard may dislodge the crawlers. 

• Monitor for continuing generations throughout 

summer. 

Pine needle scale • White, oblong scales can be found on the most 

recent year’s needles. 

• Look underneath the white scales to see if the 

purplish-red eggs have begun to hatch. 

• Paprika-colored crawlers will be visible on the 

needles. 

• Look for second generation in July. 

True fi rs Balsam woolly adelgid • Observe cut stumps for indications of the red 

reactionary wood. 



Douglas-fi r, hemlocks, 


Cryptomeria scale • Observe underside of branches for presence of 

scale. 

Elongate hemlock scale • Observe underside of branches for presence of 

scale. 

APPENDIX D .......................................................................................................................................................................................................................................................................................... 152

APPENDIX E 

INSECT TRAP USE AND 

CONSTRUCTION 

The use of traps for insect detection 

is an important method in any IPM 

program. Traps can provide an accurate 

picture of pest presence on a farm and 

may also allow for proper timing of 

pesticide applications. This proper timing 

ensures that the pesticides used will 

be effective and may ultimately lead to 

a reduction in the amount of pesticides 

used. On the following pages are 

descriptions of three insect traps that 

can be easily used in Christmas tree 

plantations. 

APPENDIX E .......................................................................................................................................................................................................................................................................................... 154

White Pine Weevil 

Detection Traps 

(Adapted from Regulatory Horticulture 

article by Sandy Gardosik and Rayanne D. 

Lehman, PDA) 

The Tedder’s trap, a baited, pyramid-shaped 

trap (Fig. 1), is used for monitoring the 

spring emergence of adult white pine weevils. 

This insect feeds on the leaders of most 

conifer trees. The dark, pyramid-shaped 

base of the trap resembles the general shape 

of a conifer, the alcohol and turpentine 

baits mimic the odors emitted by wounded 

conifers, and the funnel top captures the 

weevils much like a minnow trap. Performance 

of these traps can be greatly enhanced 

by monitoring soil temperatures at 

the trap location. (For more details, see the 

White Pine Weevil fact sheet on page 101.) 

Ordering 

The Whalon Modifi ed Tedder’s trap is sold 

by Great Lakes IPM. Each trap kit includes 

a 4-foot-tall base, a trap top, and 2 stakes. 

This trap is also advertised as a plum curculio 

and pecan weevil trap. The version sold 

as a white pine weevil trap includes 2 plastic 

vials to be used for the alcohol and turpentine 

bait. This product is listed as catalog 

# IPM-502 and sells for $16.50 in the 2010 

catalog. 

The top assembly, which consists of a 

plastic funnel and cylinder (Fig. 2), can 

be purchased separately. It is listed as the 

Boll Weevil Top Assembly, catalog # IPM- 

1006T, and sells for $1.75 (2010 catalog). 

Contact Great Lakes IPM by phone at 

1-800 235-0285 or online at www.greatlakesipm.com. 

Figure 1. 

Pyramidal 

Tedder’s trap. 

Courtesy of 

Brian Schildt, 

PDA 

Building a Trap Base 

The trap base must be prepared if only 

the trap top is purchased or if a base has 

deteriorated with usage. 

• The base consists of two triangles (see Fig. 

3 for dimensions) that may be constructed 

from any outdoor hearty material (plywood, 

corrugated plastic, etc.). A material 

with a rougher surface will provide a better 

climbing surface for the weevils. Since 

the base is intended to resemble a tree 

(to the weevils), the material should be 

dark in color—dark brown, dark green, or 

black. Paint with sand added would also 

provide a good gripping surface for the 

weevils. 

• When cutting out the triangles, cut a 

slot 24 inches long in the bottom of one 

triangle, which will be vane A. A similar 

slot 24 inches long, vane B, must be cut in 

the top of the other triangle. 

After the triangles are 

cut, holes must be drilled 

in the top of vane “A” 

(solid top, slotted 

bottom) to attach the 

bait. 

In addition, holes must 

be cut in the bottom 

of each vane, near the 

exterior angles. These 

holes will be used to 

stake the trap in place in 

the plantation. 

Figure 3. Building the trap. Courtesy of Rayanne D. Lehman, PDA 

Figure 2. 

Weevil trap top 

assembly. 

Courtesy of 

Cathy Thomas, 

PDA 

INSECT TRAP USE AND CONSTRUCTION ............................................................................................................................................................................................................................................. 155

Figure 4. Bait vials attached to 

upper portion of the trap. 


Figure 5. A kit containing baits, 

collection bags, and all other 

trap supplies. Courtesy of Cathy 

Thomas, PDA 

Figure 6. Securely wire top to 

the base. Courtesy of Cathy 

Thomas, PDA 

Figure 7. Weevils caught in trap 

(plastic vial has been removed). 


PDA 

Trap Assembly 

1. Slide triangle A (slot in bottom) over 

the top of triangle B (slot on top). The 

bottom and top edges of both triangles 

should be even. When assembled properly, 

the base will stand freely with four 

vanes. 

2. Using fi ne-gauge wire, utilize the small 

holes along the top of vane A to attach 

two small bottles. These may be the 

vials provided with the trap kit or other 

small bottles (Fig. 4). Inexpensive, 

small, glass salt and pepper shakers are a 

good option. The bottles should lie fl at 

against the angle created by the vanes 

and have their tops as close as possible 

to the top of the base. 

3. Fill one bottle with 95 percent ethyl 

alcohol (available in stores selling wine 

and spirits) or denatured alcohol (available 

in hardware stores); fi ll the other 

with gum turpentine (Fig. 5). Unless 

the bottle lids have holes, do not put 

lids on the bottles. 

4. Fit the trap top over the base, covering 

the tops of the bottles containing the 

alcohol and turpentine. Use fi ne-gauge 

wire or plastic twist ties to secure the 

top to the holes at the top of the trap 

base (Fig. 6). 

5. Twist the plastic vial over the funnel 

to screw into place. (The funnel and 

vial may come already assembled.) This 

is where the weevils will be trapped 

(Fig. 7) after crawling up the vane and 

entering through the small opening in 

the screening. 

Figure 8. Soil 

thermometer. 

Courtesy of Cathy 

Thomas, PDA 

Using the Traps 

1. Weevils become active when air temperatures 

exceed 50°F and soil temperatures 

reach 50°F (Fig. 8). Earliest 

emergence occurs in the warmest fi elds. 

For best results, the traps must be in 

place before the pests are active on a 

daily basis. Place the traps in the fi eld to 

be monitored in late winter (February) 

or early spring (early March). 

2. Locate each trap in the tree row, preferably 

next to a tree that has had weevil 

damage the previous season (Fig. 9). 

Prevent the trap from blowing over by 

using tent stakes, wire (if necessary), 

and the holes in bottoms of trap base. 

Do not use string or twine, as small 

animals will chew this and may destroy 

the base in the process. 

3. Weevils overwintering at the bases of 

trees will be attracted by the scents 

given off by the evaporation of the alcohol 

and turpentine. When they become 

active, they will climb the base of the 

trap and enter the funnel portion at the 

top. Once inside the funnel, they will 

remain inside the large plastic vial until 

removed. 

Figure 9. White pine weevil damage 

from previous season. Courtesy of 

Steven Katovich, USDA Forest Service, 


APPENDIX E .......................................................................................................................................................................................................................................................................................... 156

4. Generally, two traps per block are 

suffi cient. For extremely large blocks, 

increase the number of traps. 

5. Replenish the contents of the alcohol 

and turpentine bottles regularly. If rain 

has occurred, the bottles should be 

emptied and fi lled with fresh material. 

Comparison trials in Pennsylvania 

showed that unbaited traps did not 

capture weevils. 

6. Check the traps for weevils daily, if possible, 

during early spring. If any insects 

are in the trap, empty the contents into 

a container of soapy water or rubbing 

alcohol to kill the pests. 

7. Apply a registered pesticide to the 

terminals of susceptible trees as soon as 

possible after the fi rst white pine weevil 

is captured. During cool springs, it may 

be necessary to repeat the application if 

weevils continue to appear in the traps 

4–6 weeks after the fi rst application. 

Follow all label directions for application 

of any pesticide. 

8. These traps are for detection of weevil 

activity and will not offer control. 

Identifying the Weevils 

Several weevil species, as well as spiders, 

fl ies, stonefl ies, and other beetles, will be 

attracted to the traps (Fig. 10). It is important 

for the grower to properly identify the 

weevils that are associated with Christmas 

tree production. Four such species exist in 

Pennsylvania: white pine weevil, eastern 

(formerly northern) pine weevil, pales weevil, 

and pine root collar weevil. Both white 

pine weevil and eastern pine weevil are very 

similar in appearance. They both are dark 

red brown and have large, white and gold 

patches on the back of their wing coverings. 

To the untrained and unaided eye, the main 

difference is their size. The target species, 

white pine weevil, is the smaller of the two. 

Pales weevil and pine root collar weevil 

will also be attracted to the traps. They can 

easily be separated from the previous two 

species by size and color but are diffi cult to 

separate from each other. These weevils are 

larger and dark brown to almost black with 

small, white patches on the top of their 

wing coverings. Pine root collar weevil is 

the larger of these two species. 

Figure 10. Pales weevil (left), 

and white pine weevil (right). 

Courtesy of Gerald J. Lenhard, 

Louisiana State University, 



Figure 11. Box traps placed 

at the base of a previously 

infected tree. Courtesy of 


Figure 12. Inexpensive home 

materials used for traps. 


PDA 

Figure 13. Jar secured with duct 

tape in the box. Courtesy of 


Douglas-Fir Needle 

Midge Emergence Trap 

This home-constructed trap is a ground 

trap used for monitoring Douglas-fi r needle 

midge (Fig. 11). This insect emerges in 

spring as an adult and lays eggs in the 

newly emerging Douglas-fi r needles, where 

larvae feeding will damage the needles and 

cause them to break. The trap uses sunlight 

to attract the midge emerging from their 

overwintering spots in the soil at the base of 

Douglas-fi r trees. 

Trap Materials (Fig. 12) 

• Waxy cardboard box (approximately 

12–15 inches high and wide) 

• Clear plastic or glass jar (peanut butter jar 

works well) 

• Duct tape 

• Paper towel 

• Rocks or other weights 

Trap Assembly (Fig. 13) 

1. On one side of the box, near the bottom 

of the box (away from the opening), 

cut a hole slightly smaller than the 

mouth of the jar. 

2. Crumple the paper towel and put in 

the jar to absorb moisture and make 

detection of midges easier. 

3. Screw the jar into hole in the box and 

secure with duct tape. 

Using the Traps 

• Well before bud break, select a tree that 

had damage in previous year. If necessary, 

butt-prune the tree to allow for placement 

of the trap. 

• Place at least three traps per fi eld on the 

south side of trees. 

• Turn the box upside down so the open 

side is on the ground. Place it against the 

trunk on the southern exposure of the 

tree. 

• Secure the box in place with the rocks or 

other weights on the fl aps of box. 

• Midges that emerge from the soil under 

the box will fl y to the light/jar and collect 

there (Fig. 14). 

• As bud break approaches, monitor 

traps daily. Egg laying occurs very soon 

after emergence, so it is important that 

very little time elapses between the fi rst 

sign of midges in traps and a pesticide 

application. 

• If midges are found in the traps, empty 

the jar and destroy the midges before 

replacing the jar. 

Figure 14. Midges collected in the jar; a paper 

towel in the jar would help prevent condensation. 


APPENDIX E .......................................................................................................................................................................................................................................................................................... 158

Bagworm Traps 

Bagworm larvae feed on the foliage of many 

conifer and hardwood tree species. These 

lure-baited sticky traps (Figs. 15 and 16) 

monitor the late summer emergence of adult 

male bagworm moths. Trapping helps reduce 

the population of breeding males and, 

consequently, the number of overwintering 

eggs. 

Figure 15. 

Bagworm sticky 

trap and lure. 

Courtesy of 

Sarah Pickel, 

PDA 

Figure 16. Adult male bagworm moth (and 

other insects) caught in sticky trap. Courtesy of 


Ordering 

Bagworm traps can be ordered from Great 

Lakes IPM. The kit includes one trap top, 

one wire hanger with plastic separators, two 

sticky trap bottoms, and one lure (will last 

entire season), is listed as catalog # IPM-10, 

and sells for $3.70 (2010 catalog). 

The bagworm lure, listed as catalog 

# IPM-BGW, may be purchased separately 

for $1.55 each (2010 catalog). 

Using the Trap 

• Assemble the traps following the 

instructions provided with the trap. 

• Use two to four traps per acre in areas 

where bagworm infestations have been 

found. 

• Traps should be placed 10–15 feet away 

from the tree block being monitored. 

Hang traps 4–5 feet off the ground. 

• Begin monitoring with traps at the 

beginning of August. 

• Pesticide or Bt applications should not 

be made until the following summer and 

then only when young bagworm larvae 

are found exiting the bags. 


APPENDIX F 

SCOUTING RECORD TEMPLATE 

FOR COMMON PINE PESTS 

Regularly scouting a tree farm for pest activity is a 

major part of an integrated pest management program. 

Along with scouting, keeping accurate records 

of scouting observations is essential. When scouting 

observations are recorded each year, they can serve 

as an important reference for future growing seasons. 

The charts on the next two pages are data-recording 

tools to be used by growers when scouting in fi elds 

of various pines. These charts can be copied back to 

back and kept in a fi eld notebook, where observations 

from each scouting trip can be recorded. The 

farm/fi eld names and descriptions can be listed on 

the second chart and will serve as a reference for the 

fi rst chart. Pests common to pine species are listed 

across the chart, and below each pest several life 

stages and events are described. Growers may fi ll in 

the chart using the abbreviations given in the key 

below the chart. 

APPENDIX F ........................................................................................................................................................................................................................................................................................... 160

Scouting Record Template for Common Pine Pests 

White Pine 

Weevil 

Striped 

Pine 

Scale 

Pine Needle 

Scale 

Pine Bark 

Adelgid 

Needle Casts 

of Pines* 

Eriophyid 

Sheath Mites 

Bud 

Break 

Bagworm 

Emergence holes 

Leader wilting 

Feeding punctures/adults on leader 

Adults in trap 

Crawlers emerged 

Eggs inside female covering 

Second-generation crawlers emerged 

First-generation crawlers emerged 

Overwintering eggs 

Nymphs with white waxy fringe 

Overwintering nymphs 

Fruiting bodies/needle surface ruptured 

Yellowing of current year’s growth 

Eriophyid activity 

Eggs hatched 


Buds broken or shoots elongating (%) 

Bags present 

Larvae emerging 

Eggs present 

Farm/Field Name 

(complete reverse 

for each block) 

Date 

3/14/09 Smith/Block 4 Y 0% Y N ? Y N ? 

*Send these disease samples to PDA or Penn State for diagnosis. 

Y = Pest stage/damage found during scouting 

N = Pest stage/damage not present 

? = Did not scout for pest 

Please complete information on reverse.

Scouting Record Template for Common Pine Pests (continued) 

Please provide information for all farms and fi elds that have any species of pine planted. 

This information will be used to reference the fi eld during scouting activities listed on the front of this record. 

Farm/Field Pine Species Description (age, condition, etc.) Notes 

Smith/Field 4 Eastern white pine Planted 2004 as 2:0 from The Best Seedling Nursery History of bagworm, white pine sheath mite, and white pine weevil

APPENDIX G 


FOR COMMON SPRUCE PESTS 

Regularly scouting a tree farm for pest activity is 

a major part of an integrated pest management 

program. Along with scouting, accurate record 

keeping of scouting observations is essential. 

When scouting observations are recorded each 

year, they can serve as an important reference for 

future growing seasons. The charts on the next 

two pages are data-recording tools to be used by 

growers when scouting in fi elds of various spruces. 

These charts can be copied back to back and 

kept in a fi eld notebook, where observations from 

each scouting trip can be recorded. The farm/fi eld 

names and descriptions can be listed on the second 

chart and will serve as a reference for the fi rst 

chart. Pests common to spruce species are listed 

across the chart, and below each pest several life 

stages and events are described. Growers may fi ll in 

the chart using the abbreviations given in the key 

below the chart. 

APPENDIX G .......................................................................................................................................................................................................................................................................................... 164

Scouting Record Template for Common Spruce Pests 

White Pine 

Weevil 

Spruce 

Spider Mite 

Spruce 

Needle Rust 

Eriophyid 

Rust Mites 

Eastern Spruce 

Gall Adelgid 

Cooley Spruce 

Gall Adelgid 

Bud 

Break 

Bagworm 



Feeding punctures/ adults on leader 


>10 mites/sample unit 

Egg hatch 


Sporulation 

Symptoms 


Egg hatch 


Galls forming 



Galls forming 




Bags present 


Eggs present 

Farm/Field Name 

(complete reverse 

for each block) 

Date 





Scouting Record Template for Common Spruce Pests (continued) 

Please provide information for all farms and fi elds that have any species of spruce planted. 


Farm/Field Spruce Species Description (age, condition, etc.) Notes

APPENDIX H 


FOR COMMON TRUE FIR 

AND DOUGLAS-FIR PESTS 

Regularly scouting a tree farm for pest activity is 

a major part of an integrated pest management 

program. Along with scouting, accurate record 

keeping of scouting observations is essential. When 

scouting observations are recorded each year, they 

can serve as an important reference for future growing 

seasons. The charts on the next two pages are 

data-recording tools to be used by growers when 

scouting in fi elds of various true fi rs and Douglas-fi r. 

These charts can be copied back to back and kept 

in a fi eld notebook, where observations from each 

scouting trip can be recorded. The farm/fi eld names 

and descriptions can be listed on the second chart 

and will serve as a reference for the fi rst chart. Pests 

common to true fi rs and Douglas-fi r species are 

listed across the chart, and below each pest several 

life stages and events are described. Growers may 

fi ll in the chart using the abbreviations given in the 

key below the chart. 

APPENDIX H .......................................................................................................................................................................................................................................................................................... 168

Scouting Record Template for Common True Fir and Douglas-Fir Pests 

White Pine 

Weevil 

Spruce 

Spider Mite 

Eriophyid 

Rust Mites 

Elongate 

Hemlock 

Scale 

Cryptomeria 

Scale 

Cooley 

Spruce 

Gall 

Adelgid 

Bud 

Break 

Balsam 

Wooly 

Adelgid 

Balsam 

Twig 

Aphid 

Bagworm 



Feeding punctures/adults on leader 


>10 mites/sample unit 

Egg hatch 



Egg hatch 


Crawlers emerged 

Eggs inside female scales 

Second-generation crawlers emerged 

First-generation crawlers emerged 

Eggs observed under adult 






Stem mothers hatched 


Bags present 


Eggs present 

Farm/Field 

Name (complete 

reverse for each 

block) 

Date 





Scouting Record Template for Common True Fir and Douglas-Fir Pests (continued) 

Please provide information for all farms and fi elds that have any species of true fi r and Douglas-fi r planted. 


Farm/Field Fir Species Description (age, condition, etc.) Notes

APPENDIX I 

GROWING DEGREE DAY AND 

TEMPERATURE RECORD TEMPLATE 

Tracking growing degree days, or units of daily heat 

accumulation, is a way to pinpoint the timing of 

certain insect life cycle events—specifi cally, life stages 

when insects are vulnerable to chemical controls. 

This process involves recording daily temperatures 

and using a simple equation to convert these temperatures 

into growing degree days. The following 

chart explains the simple equation and can be copied 

and used as a record for these daily temperatures and 

growing degree day fi gures. A space for daily ground 

temperatures is also provided. This data is useful when 

monitoring for white pine weevil, a common pest 

throughout the country. Other daily weather information 

may also be useful to growers, so space is provided 

for that data. For more information on growing degree 

day and temperature monitoring, refer to page 18 of 

the IPM Basics section. 

APPENDIX I ........................................................................................................................................................................................................................................................................................... 172

Growing Degree Day and Temperature Record Template 

Remember to only record positive numbers. 

Ground temperature is helpful when monitoring for white pine weevil. 

Insert probe thermometer 2 inches into soil and look for 50°F to signal 

adult weevil emergence. 

GDD formula: 

Low Temperature + High Temperature – 50°F = GDD 

2 

Example 1: 

46°F + 60°F = 106 = 53 – 50 = 3 Add to GDD running total. 

2 2 

Example 2: 

35°F + 55°F = 90 = 45 – 50 = -5 Don’t add to GDD running total. 

2 2 

Farm: ______________________________________________________________________ Field: _____________________________________________________________________ 

Air Temperature 

Low High Avg. Daily GDD GDD Running Total Ground Temperature Other Observations (rainy, sunny, snow cover, etc.) 

Collector’s 

Initials 

Date Time

APPENDIX J 

PESTICIDE APPLICATION 

RECORD TEMPLATE 

The chart that follows is a template for growers 

to copy and record pesticide application information 

for their farms. Under the law, private 

applicators must keep records on applications 

of restricted-use pesticides. On farms where 

workers are employed, records must be kept 

on all pesticide applications, according to the 

federal Worker Protection Standard. Additionally, 

commercial/public applicators must keep 

records for every application they make. 

APPENDIX J ........................................................................................................................................................................................................................................................................................... 174

Pesticide Application Record Template 

Establishment Name: ___________________________________________________ Address: ___________________________________________________________________ 

Restricted-Use Pesticide and Worker Protection Application Records 

Application 

Rate 

EPA Reg. 

Number 

Start Time 

Applicator’s Name 

and Certifi cation 

Number 

Completion 

Time 

Quantity of 

Undiluted 

Pesticide Used 

Size of 

Treated Area 

Site, Crop, or 

Commodity 

Treated 

Active 

Ingredient 

Pesticide Brand 

Name and 

Formulation 

Application Site Location/ 

Description 

REI (hrs) 

Date 

AM 

PM AM 

PM 

AM 

PM AM 

PM 

AM 

PM AM 

PM 

AM 

PM AM 

PM 

AM 

PM AM 

PM 

AM 

PM AM 

PM

APPENDIX K 

CONTACT INFORMATION 

This appendix contains information 

pertaining to biological controls, IPM 

supplies, pest management supplies, 

and other equipment. This is an 

incomplete list, so no discrimination 

is intended and no endorsement 

by the Pennsylvania Department of 

Agriculture, Penn State Cooperative 

Extension, or Penn State’s College 

of Agricultural Sciences is implied. 

Every effort has been made to provide 

correct and up-to-date information. 

APPENDIX K .......................................................................................................................................................................................................................................................................................... 176

Association of Natural Biocontrol 

Producers (ANBP) 

PO Box 1609 

Clovis, CA 93613-1609 

Phone: 559-360-7111 

Fax: 800-553-4817 

www.anbp.org/index.htm 

The National Christmas Tree Association 

16020 Swingley Ridge Road, Suite 300 

Chesterfi eld, MO 63017 

Members/Christmas Tree Industry: 

636-449-5070 

News Media: 636-449-5071 

Fax: 636-449-5051 

E-mail: info@realchristmastrees.org 

www.christmastree.org/home.cfm 

Pennsylvania Christmas Tree Growers 

Association 

4305 North Sixth Street, Suite A 

Harrisburg, PA 17110 

Toll-free: 800-547-2842 

Phone: 717-238-9765 

Fax: 717-238-9985 

E-mail: info@christmastrees.org 

www.christmastrees.org 

Pennsylvania Landscape and Nursery 

Association (PLNA) 

1707 South Cameron Street 

Harrisburg, PA 17104-3148 

Toll-free: 800-898-3411 

Phone: 717-238-1673 

Fax: 717-238-1678 

E-mail: plna@plna.com 

www.plna.com 

Pennsylvania Department 

of Agriculture 

Sandy Gardosik 

Entomologist 

2301 North Cameron Street 


Phone: 717-772-0521 

E-mail: sgardosik@state.pa.us 

Tracey Olson 

Plant Pathologist 



Phone: 717-783-9636 

E-mail: tolson@state.pa.us 

Sarah Pickel 

Education Specialist 



717-772-5227 Phone: 

E-mail: c-sapickel@state.pa.us 

Region I Offi ce 

13410 Dunham Road 

Meadville, PA 16335-8346 

Phone: 814-332-6890 

Region II Offi ce 

542 County Farm Road, Suite 102 

Montoursville, PA 17754-9685 

Phone: 570-433-2640 

Region III Offi ce 

Route 92 South, PO Box C 

Tunkhannock, PA 18657-0318 

Phone: 717-570-2181 

Region IV Offi ce 

6 McIntyre Road 

Gibsonia, PA 15044-9644 

Phone: 724-443-1585 

Region V Offi ce 

1307 7th Street, Cricketfi eld Plaza 

Altoona, PA 16601-4701 

Phone: 814-946-7315 

Region VI Offi ce 

PO Box 5184 

Harrisburg, PA 17110-0184 

Phone: 717-346-3223 

Region VII Offi ce 

Route 113, PO Box 300 

Creamery, PA 19430-0300 

Phone: 610-489-1003 

Brian Schildt 

IPM Scouting Consultant 



Phone: 717-772-5202 

E-mail: c-brschild@state.pa.us 

Cathy Thomas 

IPM Coordinator 



Phone: 717-772-5204 

E-mail: caththomas@state.pa.us 

CONTACT INFORMATION ..................................................................................................................................................................................................................................................................... 177


Ricky Bates 

Assistant Professor of Ornamental 

Horticulture 

303 Tyson Building 

University Park, PA 16802 

Phone: 814-863-2198 

E-mail: rmb30@psu.edu 

Andrew Beck 

Extension Educator, Schuylkill and Berks 

Counties 

1202 Ag Center Drive 

Pottsville, PA 17901 

Phone: 570-622-4225 

E-mail: awb123@psu.edu 

Tom Butzler 

Extension Educator, Clinton County 

47 Cooperation Lane 

Mill Hall, PA 17751-8978 

Phone: 570-726-0022 

E-mail: tmb124@psu.edu 

Kerry Hoffman-Richards 

Acting Director 

Penn State Pesticide Education Program 

114 Buckhout Lab 


Phone: 814-865-2134 

E-mail: kmh14@psu.edu 

Greg Hoover 

Senior Extension Associate, Entomology 

543 ASI Building 


Phone: 814-865-3256 

E-mail: gah10@psu.edu 

Larry Kuhns 

Professor Emeritus of Horticulture 

304 Tyson Building 


Phone: 814-863-2197 

E-mail: ljk@psu.edu 

Robert Pollock 

Extension Educator, Indiana County 

827 Water Street 

Indiana, PA 15701-1765 

Phone: 724-465-3880 

E-mail: rcp3@psu.edu 

Suppliers 

Abraczinskas Nurseries, Inc. 

346 Numidia Drive 

Catawissa, PA 17820 

Phone: 570-356-2323 

Fax: 570-356-2366 

E-mail: info@abraczinskas.com 

www.abraczinskas.com 

B & G Equipment Company 

135 Region South Drive 

Jackson, GA 30233 

Phone: 678-688-5601 

Fax: 678-688-5633 

www.bgequip.com/index.html 

Ben Meadows Company 

PO Box 5277 

Janesville, WI 53547-5277 

Toll-free: 800-241-6401 

Fax: 800-628-2068 

www.benmeadows.com/home.htm 

Berkey’s Nursery 

138 Jefferson Street 

PO Box 215 

Spartansburg, PA 16434 

Toll-free: 800-203-9210 

Fax: 814-654-7503 

E-mail: info@berkeysnursery.com 

www.berkeysnursery.com 

BioQuip Products 

2321 Gladwick Street 

Rancho Dominguez, CA 90220 

Phone: 310-667-8800 

Fax: 310-667-8808 

www.bioquip.com 

Carl Neutzel Services, Inc. 

2648 Openshaw Road 

White Hall, MD 21161 

Toll-free: 888-580-5325 

Phone: 410-329-6791 

Fax: 410-357-4175 

E-mail: cwneutzel@verizon.net 

carlneutzel.com 

Earl F. Kegerise Inc 

3454 Pricetown Road 

Fleetwood, PA 19522 

Phone: 610-944-8532 

E-mail: larrykeg@aol.com 

Forestry Suppliers Inc. 

205 West Rankin Street 

PO Box 8397 

Jackson, MS 39284-8397 

Toll-free: 800-647-5368 

www.forestry-suppliers.com/index1.asp 

APPENDIX K .......................................................................................................................................................................................................................................................................................... 178

Gardens Alive!, Inc. 

5100 Schenley Place 

Lawrenceburg, IN 47025 

Phone: 513-354-1482 

www.gardensalive.com/default.asp 

Gempler’s 

PO Box 44993 

Madison, WI 53744-4993 

Toll-free: 800-382-8473 

E-mail: customerservice@gemplers.com 

www.gemplers.com 

Great Lakes IPM, Inc. 

10220 Church Rd NE 

Vestaburg, MI 48891 

Toll-free: 800-235-0285 

Phone: 989-268-5693 

Phone: 989-268-5911 

www.greatlakesipm.com 

The Green Spot, Ltd. 

93 Priest Road 

Nottingham, NH 03290-6204 

Phone: 603-942-8925 

Fax: 603-942-8932 

greenmethods.com/site 

Growquest Growers 

134 Davis Street 

Santa Paula, CA 93060 

Phone: 805-921-3900 

E-mail: sales@growquest.com 

www.growquest.com 

H. D. Hudson Manufacturing Company 

(Hudson Sprayers and Dusters) 

500 N. Michigan Avenue 

Chicago, IL 60611-3669 

Toll-free: 800-977-7293 

E-mail: ladybug@hdhudson.com 

www.hdhudson.com 

Hummert International 

4500 Earth City Expressway 

Earth City, MO 63045 

Toll-free: 800-325-3055 

E-mail: sales@hummert.com 

www.hummert.com 

Insects Limited, Inc. 

16950 Westfi eld Park Road 

Westfi eld, IN 46074 

Phone: 317-896-9300 

Fax: 317-867-5757 

www.insectslimited.com 

International Technology Services, Inc. 

Toll-free: 877-505-9703 

Phone: 303-661-9546 

Fax: 303-552-5747 

www.intertechserv.com 

IPM Laboratories, Inc. 

980 Main Street 

Locke, NY 13092 

Phone: 315-497-2063 

Fax: 315-497-3129 

E-mail: ipminfo@ipmlabs.com 

ipmlabs.com/home.php 

Koch’s Lawn and Garden, Inc. 

1044 Summer Valley Road 

PO Box 232 

New Ringgold, PA 17960 

Phone: 570-943-2367 

Fax: 570-943-7767 

www.kochslg.com 

Koppert Biological Systems, Inc. 

28465 Beverly Road 

Romulus, MI 48174 

Toll-free: 800-928-8827 

Fax: 734-641-3799 

E-mail: info@koppertonline.com 

www.koppertonline.com/home.asp 

Martin’s Repair Shop, LLC 

28 East Trout Run Road 

Ephrata, PA 17522 

Phone: 717-733-3015 

Fax: 717-733-9619 

E-mail: martinsreshllc@characterlink.net 

Peaceful Valley Farm Supply, Inc. 

125 Clydesdale Court 

PO Box 2209 

Grass Valley, CA 95945 

Phone: 888-784-1722 

Phone: 530-272-4769 

E-mail: helpdesk@groworganic.com 

www.groworganic.com/default.html 

Rincon-Vitova Insectaries 

PO Box 1555 

Ventura, CA 93002-1555 

Toll-free: 800-248-2847 

www.rinconvitova.com/index.htm 

Scentry Biologicals, Inc. 

610 Central Avenue 

Billings, MT 59102 

Toll-free: 800-735-5323 

Phone: 406-245-3016 

Fax: 406-245-2790 

E-mail: customerservice@scentry.com 

www.scentry.com 


Suppliers (continued) 

Sheerlund Products 

740 Corporate Drive 

Reading, PA 19605 

Toll-free: 800-233-2958 

www.sheerlundproducts.com/store/home 

.php 

Solo 

5100 Chestnut Avenue 

Newport News, VA 23605 

Phone: 757-245-4228 

Fax: 757-245-0800 

www.solousa.com/store 

Syngenta Bioline, Inc. 

PO Box 2430 

Oxnard, CA 93034-2430 

Phone: 805-986-8265 

Fax: 805-986-8267 

E-mail: Dcahn@syngentabioline.com 

www.syngenta-bioline.co.uk/Default.html 

Trécé, Inc. 

7569 Highway 28 West 

PO Box 129 

Adair, OK 74330 

Phone: 918-785-3061 

Fax: 918-785-3063 

E-mail: custserv@trece.com 

www.trece.com 

Tree-Teck, Inc. 

1773 Long Run Road 

Schuylkill Haven, PA 17972 

Toll-free: 800-477-8637 

Phone: 570-366-4997 

E-mail: info@treeteck.com 

www.treeteck.com/about.html 

Treewheeler (Vandervalk Tree Farm) 

25 Lovell Street 

Mendon, MA 01756 

Phone: 508-478-8733 

www.treewheeler.com/treewheeler.html 

APPENDIX K .......................................................................................................................................................................................................................................................................................... 180


APPENDIX L 

LISTS OF PESTS 

All of the pests that are detailed in this 

manual are listed on the following two 

pages. The fi rst list organizes the pests 

alphabetically by common name and 

includes scientifi c names and the pages of 

the corresponding fact sheets. The second 

list groups the pests by their conifer 

hosts. Several of these pests have multiple 

conifer hosts, so they may be repeated in 

the list. 

APPENDIX L ........................................................................................................................................................................................................................................................................................... 182

PESTS IN ALPHABETICAL ORDER 

Common Name Scientifi c Name Page 

Armillaria root rot Armillaria spp. 107 

Atropellis canker Atropellis tingens Lohman and Cash (also A. apiculata Lohman, Cash, and R. W. 

Davidson, A. pinicola Zeller and Goodd, and A. piniphila [Weir] Lohman and Cash) 73 

Bagworm Thyridopteryx ephemeraeformis (Haworth) 23 

Balsam twig aphid Mindarus abietinus Koch 26 

Balsam woolly adelgid Adelges piceae (Ratzeburg) 75 

Bark and engraver beetles Ips spp. and Pityogenes hopkinsi Swaine 109 

Botrytis blight Botrytis cinerea Pers.: Fr. 78 

Cooley spruce gall adelgid Adelges cooleyi (Gillette) 29, 80 

Cryptomeria scale Aspidiotus cryptomeriae Kuwana 31 

Cyclaneusma needle cast Cyclaneusma minus (Butin) DiCosmo, Peredo, and Minter 

(syn. Naemacyclus minor Butin) 34 

Diplodia (Sphaeropsis) tip blight Diplodia pinea (Desm.) J. Kickx f. 82 

Douglas-fi r needle midge Contarinia pseudotsugae Condrashoff 37 

Eastern (pine-oak) gall rust Cronartium quercuum (Berk.) Miyabe ex Shirai 88 

Eastern pine weevil Pissodes nemorensis Germar 84 

Eastern spruce gall adelgid Adelges abietis (Linnaeus) 86 

Elongate hemlock scale Fiorinia externa Ferris 39 

Eriophyid rust and sheath mites Nalepella and Setoptus spp. 42 

European pine sawfl y Neodiprion sertifer (Geoffroy) 53 

Gypsy moth Lymantria dispar (Linnaeus) 45 

Introduced pine sawfl y Diprion similis (Hartig) 53 

Japanese beetle Popillia japonica Newman 118 

Lophodermium needle cast Lophodermium seditiosum Minter, Staley, and Millar 48 

Pales weevil Hylobius pales (Herbst) 90 

Phytophthora root rot Phytophthora spp. 111 

Pine bark adelgid Pineus strobi (Hartig) 92 

Pine needle scale Chionaspis pinifoliae (Fitch) 51 

Pine root collar weevil Hylobius radicis Buchanan 113 

Pine shoot beetle Tomicus piniperda (Linnaeus) 94 

Pine wilt disease Caused by pinewood nematode, 

Bursaphelenchus xylophilus (Steiner and Buhrer) (Nickle) 116 

Ploioderma needle cast Ploioderma lethale (Dearn.) Darker 56 

Red-band (Dothistroma) needle blight Mycosphaerella pini Rostr. 

Anamorph: Dothistroma septospora (Doroguine) Morelet 58 

Redheaded pine sawfl y Neodiprion lecontei (Fitch) 53 

Rhabdocline needle cast Rhabdocline weirii A. K. Parker and J. Reid, 

Rhabdocline pseudotsugae Syd. 60 

Rhizosphaera needle cast Rhizosphaera kalkhoffi i Bubak 63 

Spruce needle rust Chrysomyxa weirii (H. S. Jacks) 65 

Spruce spider mite Oligonychus ununguis (Jacobi) 67 

Striped pine scale Toumeyella pini (King) 97 

Swiss needle cast Phaeocryptopus gäumannii (T. Rohde) Petr. 70 

Western (pine-pine) gall rust Endocronartium harknessii (J. P. Moore) Y. Hiratsuka 88 

White grubs (May, June, and Japanese beetles) Phyllophaga sp., Polyphylla sp., Popillia japonica Newman 118 

White pine blister rust Cronartium ribicola J. C. Fisch 99 

White pine weevil Pissodes strobi (Peck) 101 

Zimmerman pine moth Dioryctria zimmermani (Grote) 120 

LISTS OF PESTS ..................................................................................................................................................................................................................................................................................... 183

PESTS BY HOST GENUS 

Tree Genus Pests 

Douglas-fi r (Pseudotsuga) Armillaria root rot, bagworm, Botrytis blight, Cooley spruce gall 

adelgid, Cryptomeria scale, Diplodia (Sphaeropsis) tip blight, 

Douglas-fi r needle midge, eastern pine weevil, elongate hemlock 

scale, gyps moth, Japanese beetle, pales weevil, Phytophthora root 

rot, pine needle scale, Rhabdocline needle cast, spruce spider mite, 

Swiss needle cast, white grubs, white pine weevil, Zimmerman pine 

moth 

Fir (Abies) Armillaria root rot, bagworm, balsam twig aphid, balsam woolly 

adelgid, Botrytis blight, Cryptomeria scale, elongate hemlock scale, 

eriophyid mites, gypsy moth, Japanese beetle, pales weevil, Phytoph- 

thora root rot, pine needle scale, spruce spider mite, white grubs, 

white pine weevil 

Pines (Pinus) Armillaria root rot, Atropellis canker, bagworm, bark and engraver 

beetles, Botrytis blight, Cryptomeria scale, Cyclaneusma needle cast, 

Diplodia (Sphaeropsis) tip blight, eastern pine weevil, eriophyid mites, 

gall rusts, gypsy moth, Japanese beetle, Lophodermium needle cast, 

pales weevil, Phytophthora root rot, pine bark adelgid, pine needle 

scale, pine root collar weevil, pine sawfl ies, pine shoot beetle, Ploio- 

derma needle cast, red-band needle blight, spruce spider mite, striped 

pine scale, white grubs, white pine blister rust, white pine weevil, 

Zimmerman pine moth 

Spruce (Picea) Armillaria root rot, bagworm, Botrytis blight, Cooley spruce gall 

adelgid, Cryptomeria scale, Diplodia (Sphaeropsis) tip blight, eastern 

pine weevil, eastern spruce gall adelgid, elongate hemlock scale, 

eriophyid mites, gypsy moth, Japanese beetle, pales weevil, Phytoph- 

thora root rot, pine needle scale, Rhizosphaera needle cast, spruce 

needle rust, spruce spider mite, white grubs, white pine weevil, 

Zimmerman pine moth 

APPENDIX L ........................................................................................................................................................................................................................................................................................... 184

BIBLIOGRAPHY

Bibliography 

General Sources 

Borror, D. J., D. M. DeLong, and C. A. Triplehorn. An Introduction to the Study of Insects. 

4th ed. New York: Holt, Rinehart, and Winston, 1976. 

Chastagner, G. A, ed. Christmas Tree Diseases, Insects, and Disorders in the Pacifi c Northwest: 

Identifi cation and Management. Puyallup: Washington State University Cooperative 

Extension, 1997. 

Hansen, E. M., and K. J. Lewis. Compendium of Conifer Diseases. St. Paul: APS Press, 1997. 

Hock, W. K., and C. L. Brown, eds. Pesticide Education Manual. 2nd ed. University Park: 

The Pennsylvania State University, 1992. 

Johnson, W. T., and H. H. Lyon. Insects That Feed on Trees and Shrubs. 2nd ed. Ithaca: 

Cornell University Press, 1991. 

Kowalsick, T. “Using Growing Degree-Days for Insect Pest Management.” Ithaca: Cornell 

Cooperative Extension, 2008. ccesuffolk.org/assets/Horticulture-Leafl ets/Using-Growing- 

Degree-Days-for-Insect-Pest-Management.pdf. 

Krischik, V., and J. Davidson. “Pests of Trees and Shrubs.” In IPM of Midwest Landscapes, 

section IV. University of Minnesota, 2007. www.entomology.umn.edu/cues/IPM-trees/ 

pests_in_trees.htm. 

• “Bagworm,” pp. 71–72. Accessed on December 18, 2008. 

• “Balsam Twig Aphid,” p. 73. Accessed on December 3, 2008. 

• “Cooley Spruce Gall Adelgid,” pp. 98–99. Accessed on December 9, 2009. 

• “Eastern Spruce Gall Adelgid,” p. 108. Accessed on January 9, 2009. 

• “Eriophyid Mites,” pp. 122–23. Accessed on January 13, 2009. 

• “European Pine Sawfl ies,” p. 128. Accessed on August 28, 2009. 

• “Introduced Pine Sawfl y,” p. 155. Accessed on August 28, 2009. 

• “Pales Weevil,” p. 175. Accessed on June 9, 2009. 

• “Pine Needle Scale,” pp. 181–82. Accessed on December 3, 2008. 

• “Redheaded Pine Sawfl y,” p. 192. Accessed on August 28, 2009. 

McCullough, D. G., S. A. Katovich, M. E. Ostry, and J. Cummings-Carlson, eds. Christmas 

Tree Pest Manual. 2nd ed. East Lansing: Michigan State University Extension, 1998. 

Merrill, W., N. G. Wenner, and P. Heller. Christmas Tree Pest Notes. Department of Plant 

Pathology, College of Agricultural Sciences, The Pennsylvania State University, n.d. 

(out of print). 

Myers, T., ed. Mosby’s Medical Dictionary. 8th ed. St. Louis: Elsevier, 2009. 

Sinclair, W. A., and H. H. Lyon. Diseases of Trees and Shrubs. 2nd ed. Ithaca: Cornell 

University Press, 2005. 

Wilson, L. F. A Guide to Insect Injury of Conifers in the Lake States. Agriculture Handbook 

No. 501. Washington, D.C.: USDA Forest Service, 1977. 

IPM FOR CHRISTMAS TREE PRODUCTION ......................................................................................................................................................................................................................................... 186

Technical Sources 

Armillaria Root Rot 

“Armillaria Root Rot.” Department of Plant Pathology, Cornell University. www.plantpath 

.cornell.edu/trees/Armillaria.html (accessed on March 10, 2009). 

“Armillaria Root Rot of Trees and Shrubs.” University of Illinois Extension, Department of 

Crop Sciences. ipm.illinois.edu/diseases/rpds/602.pdf (accessed on March 10, 2009). 

Davidson, R. M., and R. S. Byther. “Armillaria (Shoestring) Root Rot (EB1776).” 

Washington State University Cooperative Extension. cru.cahe.wsu.edu/CEPublications/ 

eb1776/eb1776.html (accessed on March 10, 2009). 

Atropellis Canker 

Blosser, W. E. “Atropellis Canker of Pine.” Plant Pathology Circular No. 81. Regulatory 

Horticulture 24, no. 1 (1998). 

Lightle, P. C., and J. H. Thompson. “Atropellis Canker of Pines.” Forest Pest Leafl et 138. 

Washington, D.C.: USDA Forest Service, 1973. 

Bagworm 

Barrett, B. A. “The Bagworm in Missouri.” University of Missouri Extension. extension 

.missouri.edu/explore/agguides/pests/g07250.htm (accessed on December, 18, 2008). 

Day, E. “Bagworm.” Virginia Polytechnic Institute and State University Cooperative 

Extension. www.ext.vt.edu/departments/entomology/factsheets/bagworm.html (accessed on 

December, 18, 2008). 

Hale, F. A., B. Klingeman, and K. M. Vail. “The Bagworm and Its Control.” 

University of Tennessee Institute of Agriculture. www.utextension.utk.edu/publications/ 

spfi les/SP341-U.pdf (accessed on December 18, 2008). 

Mazzey, K., and M. Masiuk. “Woody Ornamental IPM: Bagworm.” Penn State Cooperative 

Extension. woodypests.cas.psu.edu/FactSheets/InsectFactSheets/pdf/Bagworm.pdf (accessed 

on December 18, 2008). 

Shetlar, D. J. “Bagworm and Its Control.” Ohio State University Extension Fact Sheet. 

ohioline.osu.edu/hyg-fact/2000/2149.html (accessed on December 18, 2008). 

Wheeler, A. G. Jr. “Bagworm, Thyridopteryx ephemeraeformis.” Entomology Circular No. 50. 

Regulatory Horticulture 6, no. 2 (1980): 9–12. 

Balsam Twig Aphid 

“Balsam Twig Aphid.” Center for Invasive Species and Ecosystem Health Bugwood 

Network, University of Georgia. www.forestpests.org/vermont/balsamtwig.html 

(accessed on November 20, 2008). 

Sidebottom, J. “Balsam Twig Aphid on Fraser Fir CTN-019.” North Carolina State University 

Cooperative Extension. www.ces.ncsu.edu/fl etcher/programs/xmas/ctnotes/ctn019.html 

(accessed on December 3, 2008). 

Stimmel, J. F. “Balsam Twig Aphid, Mindarus abietinus.” Entomology Circular No. 136. 


BIBLIOGRAPHY .................................................................................................................................................................................................................................................................................... 187

Technical Sources (continued) 

Balsam Woolly Adelgid 

“Balsam Woolly Adelgid.” Center for Invasive Species and Ecosystem Health Bugwood 

Network, University of Georgia. www.forestpests.org/southern/balsamwoollyadelgid.html 

(accessed on April 13, 2009). 

Day, E. “Balsam Woolly Adelgid.” Virginia Polytechnic Institute and State University 

Cooperative Extension, 1996. www.ext.vt.edu/departments/entomology/factsheets/balwoade 

.html (accessed on April 13, 2009). 

Merrill W., and N. G. Wenner. “Balsam Woolly Adelgid in Pennsylvania” Entomology 

Circular No. 128. Regulatory Horticulture 15, no. 1 (1989): 9–10. 

Ragenovich, I. R., and R. G. Mitchell. “Balsam Woolly Adelgid.” Forest Insect and Disease 

Leafl et 118. U.S. Department of Agriculture. www.fs.fed.us/r6/nr/fi d/fi dls/fi dl-118.pdf 


Sidebottom, J. “Balsam Woolly Adelgid CTN-020.” North Carolina State University 



———. “Scouting Frasier Fir in North Carolina (AG-573)—Balsam Woolly Adelgid.” 

ipm.ncsu.edu/Scouting_Fraser_Fir/contents.html (accessed on April 13, 2009). 

Botrytis Blight 

Douglas, S. M. Disease Problems in Connecticut Christmas Tree Plantations, 2004–2005. 

New Haven: Connecticut Agricultural Experiment Station, 2006. vvv.caes.state.ct.us/ 

FactSheetFiles/PlantPathology/fspp024f.htm (accessed October 7, 2009). 

———. Diseases of Christmas Tree Seedlings and Transplant Beds. New Haven: Connecticut 

Agricultural Experiment Station, 2008. www.ct.gov/caes/lib/caes/documents/publications/ 

fact_sheets/plant_pathology_and_ecology/diseases_of_christmas_tree_seedling_and_transplant_beds_06-30-08r.pdf 

(accessed October 7, 2009). 

Cooley Spruce Gall Adelgid (on Spruce and Douglas-Fir) 

“Forest Health Fact Sheet: Cooley Spruce Gall Adelgid.” Pennsylvania Department of 

Conservation and Natural Resources. www.dcnr.state.ps.us/Forestry/leafl ets/cooley.htm 


Hoover, G. A. “Cooley Spruce Gall Adelgid.” University Park: The Pennsylvania State 

University, 2002. ento.psu.edu/extension/factsheets/cooleyspgall.htm (accessed on 

December 9, 2008). 

Shetlar, D. J. “Cooley Spruce Gall Adelgid on Colorado Blue Spruce.” Ohio State 

University Extension Fact Sheet. ohioline.osu.edu/hyg-fact/2000/2552.html 


———. “Cooley Spruce Gall Adelgid on Douglas-fi r.” Ohio State University Extension 

Fact Sheet. ohioline.osu.edu/hyg-fact/2000/2551.html (accessed on December 9, 2008). 

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Cryptomeria Scale 

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Cyclaneusma Needle Cast 

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Rhizosphaera Needle Cast 

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Wood Borers and Bark Beetles 

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GLOSSARY

Glossary 

Adelgid: members of the family Phylloxeridae (Homoptera), including gall, pine, and 

woolly adelgids, that are closely related to aphids but only found feeding on conifers, some 

forming galls 

Aecium (aecia, pl.): a cup-shaped fruiting body of the rust fungi that produces aeciospores 

Aeciospores: rust fungi spores produced within aecia by infection 

Bacterium (bacteria, pl.): a single-celled, microscopic, plantlike organism that lacks 

chlorophyll and reproduces by fi ssion 

Basidiospore: an infectious spore produced by some rust fungi 

Bloom (glaucous bloom): the blue color of conifers (e.g., Colorado blue spruce) that is 

removed when sprayed with horticultural spray oil or insecticidal soap 

Brood: individuals hatching from eggs laid by one mother around the same time 

(including maturation later in life) 

Callow adults: condition of an adult shortly after emerging from the pupal case where its 

cuticle is not fully hardened or mature in color 

Cambium: a layer of actively dividing cells lying between the xylem and phloem in the 

stems and roots of vascular plants 

Candle: the growing, terminal shoot of certain conifers, particularly pines 

Canker: a disease of the bark and cambium that causes the tissue to become sunken or swollen 

Chlorosis: yellowing of normally green foliage or tissue due to chlorophyll destruction or 

failure of chlorophyll functions; often a symptom of some mineral defi ciency, extremely 

reduced light, root or stem girdling, pests, or viral infection 

Chlorotic (in terms of growth and spots): spots or splotching of normally green foliage due to 

a problem with the chlorophyll; often caused by insect feeding or disease 

Chrysalis: the pupal stage of butterfl ies; also known as the resting time between the larval 

and nymphal stages of mites 

Conidia (conidium, sing.): an asexual fungus spore 

Conidiophore: a specialized hypha bearing one or more conidia 

Contact dermatitis: a skin rash caused by an allergic reaction to something coming in 

contact with the skin (e.g., irritating hairs of gypsy moth caterpillar) 

Cull: removal of plants suspected to be infected of a disease, thus reducing the possibility of 

a spread of infection to plants not already affected 

Cuticle: with insects, the noncellular, thin, waxy outer layer of the body wall (exoskeleton) 

made up of wax and chitin; with plants, it is a thin, continuously waterproofed, noncellular 

fi lm on the external surface that tends to prevent desiccation and repels external water 

Cutin: the waxy substance comprising the inner layer of the cuticle; waterproof mixture 

of waxes, fatty acids, soaps, higher alcohols, and resinous materials forming the chief 

ingredient of the cuticle of many plants 


Desiccate (desiccation): a loss of internal moisture content necessary to maintain growth, 

rigidity (helps keep the plant up), or survival; also known as dehydration or drying 

Epidermis: with insects, the cellular layer of the body wall that secretes the cuticle; 

with plants, the outermost layer of cells occurring on all the plant parts 

Epithelial cell (epithelial cells, pl.): layer of cells lining the resin duct 

Excelsior-like strands: fi ne-curled wood shavings that result from insect feeding in the wood 

Filament: a slender, threadlike structure 

Fissure: a narrow, longitudinal opening (slit) 

Fruiting body: a fungal organ specializing in the production of spores (e.g., aecium, 

apothecia, pycnidium, tilia) 

Gallery (galleries, pl.): a tunnel or chamber made by larvae of wood-boring insects and 

composed of silk and fecal waste; usually made by bark beetles (often forming characteristic 

galleries that can be used in identifi cation) and miners and shoot, timber, and wood borers 

Host-specifi c: the degree to which an organism is limited to a specifi c type of host 

Hypha (hyphae, pl.): one of the fi laments of fungal mycelium 

Inoculum: a disease-causing propagule that can cause infection when brought into contact 

with the host 

Insect growth regulator (IGR): a pesticide built to imitate insect hormones that control 

molting and the development of some insect systems, affecting the change from immature 

to adult; in most cases, prevents the insect from becoming a sexually mature adult and may 

result in death 

Lesion: a localized area of discolored, diseased tissue resulting from an injury or wound 

Mottle (mottling): a spot or blotch of indistinct light and dark areas usually found on 

the needles 

Mycelia (mycelium, sing.): the threadlike, vegetative part of fungi; a mass of hyphae 

Necrotic: discolored or dead 

Niche (niches, pl.): a recessed compartment within a gallery where a female lays an egg 

Nuptial chamber: a large chamber created by male bark beetles used for mating with 

multiple females 

Oviposition: laying or depositing of eggs 

Ovipositor: the female insect’s egg-laying apparatus (external genitalia) 

Parasitoids: sometimes referred to as parasites and predators, these organisms feed in or on 

another living organism (host) over a long time span, consuming most to all of its tissues 

and resulting in death of the organism; behavior is intermediate between parasitism and 

predation 

Parthenogenesis (parthenogenetically): development of an egg without being fertilized 

GLOSSARY ........................................................................................................................................................................................................................................................................................... 201

Glossary (continued) 

Pathogen: a parasitic organism completely capable of causing disease 

Phloem: the layer of cells just inside bark of plants that conduct food from leaves to the 

stem and roots; different from xylem by the absence of thickened cells and the presence 

of cells containing sievelike areas 

Proleg (prolegs, pl.): a fl eshy, abdominal leg of certain insect larvae 

Pupate: larva transforming into a pupa 

Pycnidia (pycnidium, sing.): specialized, spherical, fungal fruiting bodies of rust fungi 

that produce infectious spores 

Resin duct: a tube or duct lined with epithelial cells in a woody stem or leaf, especially 

in conifers, that secretes resin (e.g., sap) 

Rhizome (rhizomes, pl.): a horizontal stem of a plant typically located underground that 

sends out roots and shoots 

Rhizomorph (rhizomorphs, pl.): rootlike structure of a fungus consisting of an aggregation 

of parallel hyphae 

Rosette: cluster of leaves in crowded circles or spirals rising up from the stem base 

(e.g., dandelion) 

Sclerotia (sclerotium, sing.): fi rm, often rounded masses of hyphae that serve as resting bodies 

and are resistant to unfavorable conditions 

Senescence: life phase of part of or a whole plant that involves decreased ability to repair 

damage, decline, or other factors that eventually lead to natural death 

Sessile: attached or fastened; incapable of moving from place to place 

Spore: single- or multiple-celled reproductive unit of fungi; each is capable of germinating 

and reproducing the organism (fungi) 

Sporulation: to produce or release spores 

Stem mother: a female aphid that produces live, identical aphids just before or as buds break 

Stippled: small dead spots on the leaves caused by mites’ and insects’ piercing-sucking 

mouthparts—specifi cally, a loss of sap and the toxic effect of saliva on the leaf tissue 

Stomata: pores in plant leaves that control gas exchange (carbon dioxide and oxygen) and 

transpiration of a plant 

Xylem: plant tissue (of trees and woody shrubs) consisting of various cell types that transport 

water and dissolved substances to the leaves; the main water-conducting tissue and chief 

supporting system composed of wood cells responsible for conduction, food transport, 

storage, and support 


Notes

Notes

Inches 

0 1 2 3 4 5 6 7 

1 

2 

3 

4 

5 

6 

7 

8 

9 

10 

Centimeters 

0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 16 17 18 19 

TAPPING SHEET FOR SCOUTING 

24 

23 

22 

21 

20 

19 

18 

17 

16 

15 

14 

13 

12 

11 

10 

9 

8 

7 

6 

5 

4 

3 

2 

1

The PA IPM Program 

is a collaboration between the 

Pennsylvania Department of Agriculture and 


aimed at promoting 

Integrated Pest Management 

in both agricultural and nonagricultural settings. 

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Produced by Ag Communications and Marketing 

© The Pennsylvania State University 2011 

Code # AGRS-117 2M11/10payne5060 

INTEGRATED 

PEST 

MANAGEMENT 

for 

CHRISTMAS 

TREE 

PRODUCTION 

A GUIDE FOR 

PENNSYLVANIA 

GROWERS

Integrated Pest Management for Christmas Tree Production: A ...

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