New Pest Response Guidelines

Cydalima perspectalis

Box tree moth

Box tree moth adult (Source: Wim Rubers at waarneming.nl, distributed under a CC-BY 3.0 license)
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Contents

Figures and Tables ............................................................................................... 4

1. Introduction ................................................................................................... 6

2. Pest Overview ................................................................................................ 7

3. Pest Identification ........................................................................................ 12

4. Delimitation Survey ..................................................................................... 19

5. Eradication and Control Options .............................................................. 28

Literature Cited ................................................................................................. 31

Appendix A: Environmental Compliance ...................................................... 37

Appendix B: Sampling ....................................................................................... 41

Appendix C: Plastic Bucket Trap Protocol ..................................................... 43

Authors and Reviewers ...................................................................................... 53
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Figures and Tables

Figures

Cover Box tree moth adult......................................................................... 1

Figure 3-1 Adult male box tree moth ............................................................ 13

Figure 3-2 Adult male brown box tree moth ................................................. 14

Figure 3-3 Eggs of box tree moth on boxwood leaf ...................................... 14

Figure 3-4 Newly hatched box tree moth larvae ........................................... 15

Figure 3-5 Box tree moth larvae .................................................................... 15

Figure 3-6 (A and B) Box tree moth pupae surrounded by white silk and host 

foliage; (C and D) pupae exposed after removal of white silk and 

host foliage ................................................................................... 16

Figure 3-7 Mature box tree moth larva feeding on leaves of common 

boxwood ....................................................................................... 17

Figure 3-8 Box tree moth damage to common boxwood .............................. 17

Figure 3-9 Common boxwood defoliation and webbing due to box tree moth 

feeding .......................................................................................... 18

Figure 3-10 Adult melonworm ........................................................................ 18

Figure 3-11 Boxwood dieback of twigs (A) with dying leaves, and (B) black 

discoloration of stem under bark................................................... 19

Figure 4-1 Standard core infested area after an initial detection ................... 21

Figure 4-2 Alternate core infested area size .................................................. 22

Figure 4-3 Trapping transects extending out from a core infested area in the 

cardinal directions......................................................................... 23

Figure 4-4 Additional trapping transects extending out from a core infested 

area in the ordinal directions......................................................... 23

Figure 4-5 Placing traps in a core infested area with a 0.5 mi radius............. 24

Figure 4-6 Placing traps along a trapping transect ......................................... 25
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Figure 4-7 Survey expansion after detecting moths in the core 

infested area .................................................................................. 27

Figure 4-8 Survey expansion after detecting moths in the trapping 

transects......................................................................................... 28

Tables

Table 2-1 Duration of box tree moth life stages and intervals under laboratory 

conditions at 77 °F .......................................................................... 9

Table 2-2 Box tree moth hosts ...................................................................... 10

Table 5-1 Insecticides available in the United States for use against box 

tree moth ....................................................................................... 30
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1. Introduction

Plant Protection and Quarantine (PPQ) develops New Pest Response Guidelines 

(NPRGs) in preparation for potential pest introductions. This document is based 

on the best information available at the time of development and may not reflect 

the latest state of knowledge at the time the pest is detected. In addition, the PPQ 

response must be tailored to the specific circumstances of each pest introduction 

event, which cannot be predicted. Therefore, this document provides only general 

guidelines to be used as a basis for developing a situation-specific response plan 

at the time a new pest is detected.

Program managers of Federal emergency response or domestic pest control 

programs must ensure that their programs comply with all Federal Acts and 

Executive Orders pertaining to the environment, as applicable. Refer to the 

Environmental Compliance section in Appendix A for details.

Chapter

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2. Pest Overview

Key Information

♦ Box tree moth, Cydalima perspectalis, is native to East Asia and has 

become a serious invasive pest in Europe, where it continues to spread. In 

2018, it was found in Ontario, Canada. 

♦ The box tree moth feeds primarily on boxwood (Buxus spp.).

♦ Larvae feed on leaves, and heavy infestations can defoliate host plants. 

Once the leaves are gone, larvae consume the bark, leading to girdling and 

plant death.

♦ In introduced areas where box tree moth has 2 generations per year, 

boxwood stands have declined over 95% in 8 years or less.

♦ Long-distance spread of the box tree moth is mainly through movement of 

infested boxwood.

♦ The box tree moth can be identified based on morphological 

characteristics. 

♦ The most effective way to survey for box tree moths is with bucket traps 

with pheromone lures.

♦ Box tree moth is difficult to eradicate once established but can be

effectively controlled with chemical sprays and Bacillus thuringiensis 

kurstaki (Btk). 

Taxonomy

Scientific Name

♦ Cydalima perspectalis (Walker)

Taxonomic Position

♦ Animalia: Arthropoda: Insecta: Lepidoptera: Crambidae

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Synonym(s)

♦ Diaphania perspectalis (Walker)

♦ Glyphodes perspectalis (Walker)

Common Name(s)

♦ Box tree moth

♦ Box tree pyralid

Biology and Ecology

The box tree moth is native to temperate and sub-tropical regions in Asia (Mally 

and Nuss, 2010). It was first reported in Europe in 2007, after which it spread 

rapidly across Europe into Western Asia and Northern Africa (Agius, 2018; Geci 

et al., 2020; Haddad et al., 2020; Hizal et al., 2012; Leuthardt and Baur, 2013; 

Perez and Guillem, 2019). In 2018, it was documented in Canada (Plant et al., 

2019).

Life Cycle

Developmental times for the box tree moth vary with temperature. At 77 °F, the 

total life cycle (from eggs hatching to adults laying eggs) is between 33 and 44 

days (See Table 2.1). Two to five generations may occur per year, depending on 

climatic conditions (Wan et al., 2014).

Eggs

Females lay eggs singly or in clusters of 5 to more than 20 eggs in a gelatinous 

mass on boxwood leaves (See Species ID/Diagnostic) (Farahani et al., 2021; 

Leuthardt and Baur, 2013). Most females deposit more than 42 egg masses in 

their lifetime (Leuthardt and Baur, 2013). Temperature determines the 

development time of the eggs (Stan and Mitrea, 2020), but they typically hatch 

within 4 to 6 days (Farahani et al., 2021).

Larvae

Box tree moths overwinter as larvae, and all six larval stages are capable of 

diapausing (Maruyama and Shinkaji, 1991; Nacambo et al., 2014). Overwintering 

larvae emerge in response to rising temperatures (Poitou et al., 2020). While the 

timing varies based on local climate, larvae typically begin feeding in March, 

continuing until they pupate in late April to early May (Farahani et al., 2021; 

Nacambo et al., 2014; Stan and Mitrea, 2020).
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Larvae of the next generation hatch in May or June and immediately begin 

feeding on boxwood leaves. As they develop, they spin silken webs to hold leaves 

together and create protected areas to feed (Matošević, 2013; Nacambo et al., 

2014). They tend to feed on leaves in the lower portion of host plants, but reside 

in the upper portion (Kulfan et al., 2020). Larvae remain active through the 

summer until September or October. Then, in response to shortened daylight 

hours, larvae will construct a cocoon between two leaves and enter diapause 

(Xiao et al., 2011). 

Pupae

Pupation occurs on the host leaves in silk cocoons (Korycinska and Eyre, 2011). 

If the boxwood host is defoliated, pupation may occur away from the host plant 

using leaves from the surrounding area (Bras, 2021). Pupae will typically first 

appear in April or May and will be present continuously through the summer and 

into the fall, depending on the local climate and timing of generations. See Table 

2.1 for pupal duration. 

Adults

Adult females use sex pheromones to attract mates (Kim and Park, 2013). They 

are highly mobile (See Dispersal) and are typically active at dusk and throughout 

the night (Székely et al., 2011). 

Adults first emerge from the overwintering generation between April and July, 

depending on climate and temperature. Subsequent generations are active between 

June and October (Chen et al., 2005; Farahani et al., 2021; Göttig and Herz, 2017; 

Nacambo et al., 2014). Adults typically live for two weeks after emergence (Bras, 

2021). See Table 2.1 for adult lifespan range. 

Table 2-1 Duration of box tree moth life stages and intervals under laboratory 

conditions at 77 °F (Farahani et al., 2021) 

Life stage/interval Duration (minimummaximum) in days

Egg 4–6

1st instar larvae 2–5

2nd instar larvae 2–7

3rd instar larvae 2–4

4th instar larvae 2–5

5th instar larvae 3–6

6th instar larvae 5–12

Prepupae 1–2

Total larval period 20–31

Pupae 6–8

Time from Egg to Adult 33–44

Female adult life span 5–27
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Life stage/interval Duration (minimummaximum) in days

Male adult life span 4–25

Total life span 39–62

Hosts1

The box tree moth completes its development and causes extensive damage 

mainly on boxwood (Buxus) species (John and Schumacher, 2013; Leuthardt and 

Baur, 2013; Matsiakh et al., 2018; Tuniyev et al., 2016). It has also been shown to 

complete its development on Murraya paniculata (Wang, 2008). 

The only other plant species that may support box tree moth development, based 

on current knowledge, are Euonymus alatus and E. japonicus, though the 

available evidence is not conclusive (PPRA, 2021). For Ilex purpurea or any other 

plants on which feeding has been reported in the literature, there is currently no 

convincing evidence that they support box tree moth development (PPRA, 2021). 

See Table 2-2 for box tree moth hosts. 

Table 2-2 Box tree moth hosts 

Family Scientific 

name

Common 

name

Notes and References Confidence

Buxaceae Buxus 

microphylla

littleleaf 

boxwood

Maruyama, 1992 Host status 

certain

Buxaceae Buxus 

microphylla

var. japonica

Japanese 

boxwood

Maruyama, 1992 Host status 

certain

Buxaceae Buxus 

sempervirens

common 

boxwood

Hizal, 2012 Host status 

certain

Buxaceae Buxus sinica

var. insularis

(=Buxus

microphylla

var. insularis

Korean 

boxwood

Maruyama, 1992 Host status 

certain

Rutaceae Murraya 

paniculata

mock 

orange

Wang, 2008 Host status 

certain

Celastraceae Euonymus 

alatus

burningbush Larvae are documented feeding 

on this plant in the field (Chen 

and Chen, 2017; Tu et al., 2017; 

Zhang and Wang, 2012), but 

laboratory results are mixed. Two 

laboratory tests showed minimal 

feeding and no development

(Brua, 2013; Wiesner et al., 

2021). One report stated adults 

were reared from larvae that 

were fed this plant, but no details 

were provided (Brua, 2013).

Host status 

uncertain

1 Hosts are plants that support box tree moth development.
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Family Scientific 

name

Common 

name

Notes and References Confidence

Celastraceae Euonymus 

japonicus

Japanese 

spindletree

Most reports refer to this plant 

using a Chinese common name 

that is also used for Buxus 

megistophylla (Chen, 2008; Liu et 

al., 2009; Ma and Yang, 2010; Qi, 

2003; Sun et al., 2009; Wang et 

al., 2013). 

Larvae are documented feeding 

on this plant in the field (Chen, 

2008; Gu and Yang, 2006; Liu et 

al., 2009; Ma and Yang, 2010; Qi, 

2003; Sun et al., 2009; Wang et 

al., 2013). Laboratory studies, 

however, have shown that this 

plant does not support complete 

development (Brua, 2013; 

Karpun et al., 2015; Matošević et 

al., 2017; van der Straten and 

Muus, 2010).

Host status 

uncertain

Dispersal

The rate of spread for the box tree moth has varied since its introduction to 

Europe, with some cases peaking at 96 miles per year (Roques et al., 2016).

Human-Assisted Spread

Long distance movement of the box tree moth across Europe occurred through the 

movement of infested boxwood plantings (Bras et al., 2019). Multiple detections 

have been made at nurseries where boxwoods are commonly grown and shipped 

over long distances (Bella, 2013). 

Natural Dispersal

Box tree moths are highly mobile and are reported to be good fliers (Bella, 2013).

Natural spread of this moth in Europe is about 3-6 miles per year (John and 

Schumacher, 2013; van der Straten and Muus, 2010). One analysis from Europe 

concluded that natural dispersal from continental Europe to the United Kingdom 

was possible, suggesting sustained adult flights of over 20 miles (Korycinska and 

Eyre, 2011).
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3. Pest Identification 

Species ID/Diagnostic

Morphological

Adults

Bodies are white, with a brown head and abdomen (Fig. 3-1) (Matošević, 2013). 

Wings are white and slightly iridescent, with an irregular thick brown border

spanning 1.6–1.8 inches (Gutue et al., 2014; Korycinska and Eyre, 2011). Some 

adults have completely brown wings with a small white streak on each forewing 

(Fig. 3-2) (Albert, 2009; Korycinska and Eyre, 2011). Males and females show 

both colorations (Bras, 2021).

Figure 3-1 Adult male box tree moth (Source: Szabolcs Sáfián, Bugwood.org)

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Figure 3-2 Adult male brown box tree moth (Source: Szabolcs Sáfián, Bugwood.org)

Eggs

Eggs are pale yellow, averaging 0.04 inches and laid in flat clusters (Fig. 3-3)

(Hizal et al., 2012; Korycinska and Eyre, 2011). As they mature, a black spot 

appears marking the larval head (Korycinska and Eyre, 2011).

Figure 3-3 Eggs of box tree moth on a boxwood leaf (Source: W. Schön, 

www.schmetterling-raupe.de)
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Larvae

Newly hatched larvae have black heads and are green to yellow (Fig. 3-4). As 

they age, dark brown stripes develop on the body. 6th instar larvae are about 1.6 

inches long and have thin white and thick black stripes and black dots outlined in 

white along the length of the body (Fig. 3-5) (Korycinska and Eyre, 2011).

Figure 3-4 Newly hatched box tree moth larvae (Source: Colette Walter, 

http://www.schmetterling-raupe.de/art/perspectalis.htm)

Figure 3-5 Box tree moth larvae (Source: Böhringer Friedrich, at rufre@lenz-nenning.at, 

distributed under a CC-BY 3.0 license)
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Pupae

Pupae develop inside a silk cocoon and are 0.6–0.8 inches long. They are initially 

green, with black stripes on the back, and turn brown as they mature (Fig. 3-6)

(Korycinska and Eyre, 2011).

Figure 3-6 (A and B) Box tree moth pupae surrounded by white silk and host foliage; (C 

and D) pupae exposed after removal of white silk and host foliage (Sources: Korycinska 

and Eyre, 2011 and http://www.schmetterling-raupe.de/art/perspectalis.htm)

Signs and Symptoms

Signs of damage may not appear at the beginning of an infestation because young 

larvae hide among twigs and leaves (Hrnčić and Radonjić, 2017). Larvae 

skeletonize the leaves and feed on the bark, causing defoliation and dryness, 

leading to the host’s death (Figs. 3-7 and 3-8) (Hrnčić and Radonjić, 2017; 

Nacambo et al., 2014). Signs of feeding include green-black frass and webbing 

(Fig. 3-9) (Gutue et al., 2014). 
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Figure 3-7 Mature box tree moth larva feeding on leaves of common boxwood (Source: 

Ferenc Lakatos, University of West-Hungary, Bugwood.org)

Figure 3-8 Box tree moth damage to common boxwood (Source: Ferenc Lakatos, 

University of West-Hungary, Bugwood.org)
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Figure 3-9 Common boxwood defoliation and webbing due to box tree moth feeding 

(Source: Colette Walter, http://www.lepiforum.de/webbbs/images/forum_2/pic13983.jpg)

Similar Species

The melonworm, Diaphania hyalinata, is found in the United States in south 

Florida and possibly south Texas and feeds on the leaves of cucurbits (Fig. 3-10)

(Capinera, 2020). The adult moth is similar in appearance to the box tree moth.

Figure 3-10 Adult melonworm (Source: Jack Foreman, distributed under a CC-BY 3.0 

license)
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Boxwood dieback caused by the fungal pathogen, Colletotrichum theobromicola, 

causes dieback of twigs with light tan leaves that remain attached to the branches

(Singh and Doyle, 2017). The infection also causes a black discoloration under 

the bark of twigs (Singh and Doyle, 2017). This disease of boxwoods has been 

found in Louisiana, North Carolina, South Carolina and Virginia (Singh et al., 

2015).

Figure 3-11 Boxwood dieback of twigs (A) with dying leaves, and (B) black 

discoloration of stem under bark (Source: Raj Singh, LSU AgCenter)
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4. Delimitation Survey

Delimitation Area

Delimitation surveys determine the extent of the infested area after a pest has 

been found. The size and shape of the delimitation area may differ from one 

location to another due to differences in host plant density and distribution, 

topography, and agency resources at the time of pest introduction. 

Timing of Surveys

Start the delimitation surveys immediately after detecting box tree moth. 

Depending on the climatic conditions where the moth is detected, two to five 

generations may occur per year. If no additional detections are made, surveys 

should continue for at least one year in areas with three or more generations and 

for two years in areas with two generations. 

Survey continuously when temperatures are greater than 51°F and adults are 

flying, typically between May and October. If moths are detected in the fall in 

areas where the pest is likely to overwinter, surveys should resume the following 

spring, based on degree day models for adult emergence.

Type of Survey

The most suitable method for detecting and delimiting box tree moth is through 

the use of pheromone traps. The survey design focuses on trapping in the core 

area around the initial detection and along the perimeter of the survey area. 

Visual inspections for box tree moths and damage to host plants (see Signs and 

Symptoms for descriptions) can be useful, but some infested plants may not have

visible insects or symptoms, depending on the time and severity of infestation.

Chapter

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Delimitation Survey Design

To delimit box tree moth using a core and transect design:

1. Identify the core infested area and the extended survey transects.

a. Using a map or mapping software, draw a circle that extends out from 

the initial detection (Fig. 4-1).

i. Standard surveys should have a core infested area extending 0.5 

miles from the initial detection.

ii. Under certain scenarios, a smaller (0.3 mile radius) or larger (1 

mi radius) core infested area will be adequate (Fig. 4-2).

1. Use a smaller core if resources are limited or traps are 

not available and a larger core if enough resources are 

available. 

2. Use a smaller core if the detection is made where moths 

are unlikely to disperse (e.g., in a nursery where there 

are plentiful host plants). 

3. Use a larger core if the pest population is wellestablished near the initial detection and is more likely 

to have dispersed long distances.

Figure 4-1 Standard core infested area after an initial detection
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Figure 4-2 Alternate core infested area sizes

b. Using a map or mapping software, draw four transect lines.

i. If moths are detected at a retailer or nursery, surveyors may use 

either the retailer’s service area if that information is available 

to determine the length and direction of surveying transects.

ii. If there isn’t any information to suggest a direction for transects, 

place them in each cardinal direction (Fig. 4-3).

iii. If more traps are available and if there is no information 

available regarding the extent or direction of spread, consider 

drawing four additional transect lines in each of the ordinal 

directions (NE, SE, NW, SW) (Fig. 4-4). 

iv. Standard transect length should be 3 miles from the initial 

detection. 

1. A standard core area with radius of 0.5 miles will have 

2.5-mile transects.

2. A smaller core area (0.3-mile radius) will have 2.7-mile 

transects and a larger core (1-mi radius) will have 2-

mile transects.
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Figure 4-3 Trapping transects extending out from a core infested area in the cardinal 

directions

Figure 4-4 Additional trapping transects extending out from a core infested area in 

the ordinal directions
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2. Survey the core infested area.

a. Place traps throughout the core infested area at a density of 64 traps 

per square mile2

.

b. A core infested area with a radius of 0.5 miles covers 0.784 square 

miles, which means you will need at least 50 traps to survey this area 

(Fig. 4-5). A 0.3-mile radius will require at least 18 traps and a 1-mile 

radius will require at least 201 traps. 

c. Space traps regularly throughout the area, at least 66 feet apart, and 

near hosts when possible. See the Sampling Instructions for details.

d. Traps should be checked at least every two weeks and should be kept 

in place according to the guidelines in Timing of Surveys. 

Figure 4-5 Placing traps in a core infested area with a 0.5 mi radius. Trapping 

density is 64 traps per square mile. Traps needed will change with a smaller or larger 

core.

3. Survey along the transect lines (Fig. 4-6). 

a. Each 2.5 mi transect requires up to 15 traps, based on the trap-to-trap 

distance when using 36 traps per square mile. Consult the survey 

design tool (included separately) to adjust the trap placement to fit the 

2 Consult the survey design tool (included separately) to see the probabilities of detection for 

different trap densities.
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needs of the survey.

b. For a 2.5 mi transect with 36 traps per linear mile, place traps at least 

866 feet apart. Consult the survey design tool (included separately) to 

adjust the trap spacing with the change in trap density. 

c. Only place traps in areas with available hosts.

d. Barren areas or non-permeable surfaces should be excluded. Do not 

place traps in these areas.

e. If areas with continuous hosts and suitable habitats are available along 

the entire transect, surveyors will deploy 15 traps spaced evenly. 

f. Up to 60 traps are required to fully cover transects in each cardinal 

direction (Fig. 4-3). Up to 120 traps are required to fully cover 

transects placed in both the cardinal and ordinal directions (Fig. 4-4). 

g. Follow the Sampling Instructions for details on trap placement.

h. Traps should checked at least every two weeks and should be kept in 

place according to the guidelines in Timing of Surveys.

Figure 4-6 Placing traps along a trapping transect. Place traps in habitat with hosts 

and avoid placing traps in unsuitable areas. If suitable areas are continuous along 

the 2.5 mile transect, 15 traps will be used.

4. Employ an Outreach Program to raise awareness and detect box tree moth 

over larger areas where surveys are not feasible.
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Survey Adjustments and Expansion

Adjustments

As data become available after traps in the field are serviced, surveyors may need 

to adjust their trapping protocols. If there are no detections in the transect, 

consider changing the location of the transects to cover more area outside the 

core. If there are no detections in the core, continue trapping in the core for about 

three months to capture potential next generation adults. After confirming that 

there are no new populations in the core, the traps may be moved to areas outside 

the core. If moths are found, surveyors can adjust the trapping density in the core 

to focus on specific locations or add/extend trapping transects to better delimit 

pests near additional detections outside the core. 

To more effectively place additional traps, consider investigating whether moths 

found in the core or along trapping transects originated from nearby areas. 

Surveyors may look for infested hosts around the trap or at high-risk sites such as 

retailers or nurseries in the vicinity of the trap (see Signs and Symptoms for 

images of infestations). If any infested hosts are found, surveyors can adjust 

trapping locations or density based on the source of infestation or direction of 

spread, as determined by trace-back and trace-forward surveys.

Survey Expansion in the Core Area

If any new detections are made within 0.3 miles of initial detection, do not expand 

the survey. Continue trapping in the area, following the guidelines from Timing of 

Surveys. Detections close to the initial detection will inform control efforts by 

determining pest density and dispersal direction. 

If any detections are made between 0.3 miles and 0.5 miles of the initial detection, 

expand the core from the point of additional detection in a circle with a radius of 

0.3 miles (Fig 4-7). Place traps throughout the expanded core infested area at a 

density of 64 traps per square mile. In addition, add an ordinal trapping transect 

(NE, NW, SE, SW) in the quadrant where the detection was made, if ordinal 

transects have not already been placed.
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Figure 4-7 Survey expansion after detecting moths in the core infested area. 

Surveyors should add an ordinal transect if one has not already been included in the 

survey.

Along the Trapping Transects

Additional detections along trapping transects may indicate a new focal point of 

the infestation and require survey expansion. After any detection from a trapping 

transect, use mapping software to create an additional core infested area 

(extending 0.5 miles from the new detection) and a new set of trapping transects 

extending 3 miles from the detection).

Surveyors may need to adjust the size of the core, the length of the trapping 

transects, and the direction of the trapping transects based on the available moth 

distribution data. Do not place additional traps in previously surveyed areas. If the 

detection is relatively close to the initial detection, placing transects at different 

angles will help cover unsurveyed areas (Fig. 4-8).
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Figure 4-8 Survey expansion after detecting moths in the trapping transects. 

Expanded trapping transects are placed at 45 degree angles to the original transects 

to better cover the area.

Outreach Program for Box Tree Moth

Developing an outreach campaign as a supplement for traditional surveys will 

more effectively detect box tree moth over a large area. This outreach should be 

organized and carried out by state extension services and operate side by side with 

traditional detection surveys by PPQ staff. This campaign should target surveyed 

and at-risk areas near the initial detection. It should encourage nurseries and 

landscape managers in the region to proactively inspect hosts, place sentinel traps 

around hosts, and report infestations (see Signs and Symptoms) to the proper 

authorities. Photographs of box tree moths on plants should be submitted to local 

extension agents or PPQ staff to help them identify new detections and areas for 

further investigation. 
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5. Eradication and Control 

Options

Overview

This information can be used by PPQ decision-makers after a detection to assess 

the suitability of potential actions to eradicate, contain, or suppress box tree moth. 

Although eradication should always be prioritized, its success will depend on the 

pest population size and distribution. Use the chemical and Bt-based options listed 

below to attempt eradication of the pest.

Eradication Options

There is no evidence of successful eradication of the box tree moth where it has 

been introduced (CABI/EPPO, 2018). Eradication of established populations in 

Europe, including in Germany and Switzerland, is no longer considered feasible 

(Albert, 2009; van der Straten and Muus, 2010). However, eradication is possible 

if the pest is contained soon after introduction in smaller patches of cultivated or 

highly managed boxwoods. In these cases, chemical sprays should be used to kill 

larvae on infested hosts. In areas where the moth has become widely established, 

use the control options listed below to contain or suppress the box tree moth. 

Control Options

Host Removal

If heavily infested boxwoods are visually detected, remove and destroy the 

infested shrubs and any nearby hosts to prevent further rapid spread of the moth.

♦ Hosts can be destroyed by burial, chipping, or burning. In case of burning, 

check local ordinances for guidelines and required documentation.

♦ Treat uninfested and unremoved hosts prophylactically with a pyrethroid 

or organophosphate (see Chemical Control, below).

Be sure to contain affected plants to safeguard the material while it is being 

Chapter

5
Last update 02FEB2022 Cydalima perspectalis 29

moved to another site for destruction, especially when chemical control is not 

feasible. Placing plant material (e.g. potted or removed plants) in large plastic

bags may prevent insect escapes. Placing bagged material in the sun will increase 

temperatures that may be lethal to the insect. 

Chemical Control

Chemical controls may require high spray volume for effective control of box tree 

moth since the caterpillars feed on the undersides of leaves and create webbing 

that may protect them from adequate exposure. In Asia, pyrethroids (deltamethrin, 

cypermethrin), organophosphates (chlorfluazuron), spinosyns (spinosad), and 

phenylpyrazoles (fipronil) are used against box tree moth (Wan et al., 2014). 

Spinosad and fipronil are recommended treatments in parts of China (Wan et al., 

2014) and are most effective against early larval stages (Kenis, 2016). In infested 

areas of Europe, chemical treatments have been the primary means to protect 

Buxus plantings (Kenis et al., 2013). Table 5-1 displays the most effective active 

ingredients tested against box tree moth larvae. Insecticide options are not 

available to use on pupae. 

Table 5-1 Insecticides available in the United States for use against box tree moth

IRAC Insecticide Class 

(Mode of Action)1

Active Ingredient Efficacy Registered 

in the U.S.*

References for use

Diamides (28) Chlorantraniliprole 99% yes Somsai et al., 2019

Ecdysone Receptor

Antagonists (18)

Methoxyfenozide 99% yes Somsai et al., 2019

Glutamate gated 

chloride channel blocker

(GluCl) (6)

Abamectin 100% yes** Somsai et al., 2019

Glutamate gated 

chloride channel blocker

(GluCl) (6)

Emamectin 

benzoate

>95% yes Qian et al., 2018

Neonicotinoids (4A) Thiacloprid 96% yes*** Stan and Mitrea, 2019

Organophosphate (1B) Trichlorfon >95% yes** Qian et al., 2018

Organophosphate (1B) Dimethoate 100% yes Raspudić et al., 2018

Organophosphate (1B) Chlorpyrifos 93% yes Somsai et al., 2019

Pyrethroids (3A) λ-cyhalothrin 98% yes Stan and Mitrea, 2019

Pyrethroids (3A) β-cyfluthrin >95% yes Qian et al., 2018

Pyrethroids (3A) Cypermethrin 98% yes Stan and Mitrea, 2019

Pyrethroids (3A) τ-Fluvalinate 98% yes** Stan and Mitrea, 2019

Spinosyns (5) Spinosad 99% yes Somsai et al., 2019

1 MOA: Insecticide acts on nerve & muscle (1B, 3A, 4A, 5, 6, 28), or acts as an insect growth regulator (18). For full definitions

for each mode of action see https://irac-online.org/modes-of-action. 

*Registration information collected from CDMS, 2021. No active ingredients are labeled specifically for box tree moth. 

**Registered in the United States, but not typically labeled for lepidopteran pests. 

***Registered in the United States, with no commercial products available. 
Last update 02FEB2022 Cydalima perspectalis 30

Biological Controls

Egg

Chelonus tabonus, Tyndarichus spp. and Trichogramma spp. are reported to 

parasitize box tree moth eggs (Göttig and Herz, 2016; Wan et al., 2014). 

Larva

Bioinsecticides based on Bacillus thuringiensis serotype kurstaki (Btk) have been 

effective against box tree moth larvae in Europe (Guérin, 2018) and are now the 

recommended control method in that region (Göttig and Herz, 2018; Lefort et al., 

2014).

Casinaria spp., Compsilura concinnata, Dolichogenidea stantoni, Exorista sp., 

Protapanteles mygdonia and Pseudoperichaeta nigrolineata are larval parasitoids 

of box tree moth (Belokobylskij and Gninenko, 2016; Farahani et al., 2018; Shi 

and Hu, 2007; Wan et al., 2014). In laboratory studies, entomopathogenic 

nematodes Steinernema carpocapsae and Heterorhabditis bacteriophora caused 

high mortality of box tree moth larvae. 

Pupa

Brachymeria lasus and Apechthis compunctator are pupal parasitoids of box tree 

moth (Wan et al., 2014). 

Cultural Controls

Removing larvae manually or via water-spraying has been recommended for 

control on ornamental box trees, but not for forests or other large patches of 

unmanaged hosts (Alkan Akıncı and Kurdoğlu, 2019; Wan et al., 2014). Manual 

removal is labor intensive, and water-spraying is difficult and may damage the 

tree (Kenis, 2016). Recommendations for parks and gardens in China include

visual inspection and removal of eggs every two to three days during the 

oviposition season (Shi and Hu, 2007).
Last update 02FEB2022 Cydalima perspectalis 31

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Environmental Compliance

Introduction

Use Appendix A as a guide to environmental regulations pertinent to the box tree 

moth.

Overview

Program managers of Federal emergency response or domestic pest control 

programs must ensure that their programs comply with all Federal Acts and 

Executive Orders pertaining to the environment, as applicable. Two primary 

Federal Acts, the National Environmental Policy Act (NEPA) and the Endangered 

Species Act (ESA), often require the development of significant documentation 

before program actions may commence. Environmental and Risk Analysis 

Services (ERAS), a unit of APHIS’ Policy and Program Development Staff 

(PPD), is available to provide guidance and advice to program managers and 

prepare drafts of applicable environmental documentation. In preparing draft 

NEPA documentation ERAS may also perform and incorporate assessments that 

pertain to other Acts and Executive Orders, described below, as part of the NEPA

process. The Environmental Compliance Team (ECT), a part of PPQ’s Plant 

Health Programs (PHP), assists ERAS in development of documents and 

implements any environmental monitoring. Program leadership is strongly 

advised to consult with ERAS and/or ECT early in the development of a program 

in order to conduct a preliminary review of applicable environmental statutes and 

to ensure timely compliance. 

Environmental monitoring of APHIS pest control activities may be required as 

part of compliance with environmental statutes, as requested by program 

managers, or as suggested to address concerns with controversial activities.

Monitoring may be conducted with regards to worker exposure, pesticide quality 

assurance and control, off-site chemical deposition, or program efficacy. Different 

tools and techniques are used depending on the monitoring goals and control 

techniques used in the program. Staff from ECT will work with the program 

manager to develop an environmental monitoring plan, conduct training to 

implement the plan, provide day-to-day guidance on monitoring, and provide an 

Appendix

A
Last update 02FEB2022 Cydalima perspectalis 38

interpretive report of monitoring activities.

The following is list of pertinent laws and Executive Orders:

National Environmental Policy Act (NEPA) – NEPA requires all Federal 

agencies to examine whether their actions may significantly affect the quality of 

the human environment. The purpose of NEPA is to inform the decision-maker 

prior to taking action and to inform the public of the decision. Actions that are 

excluded from this examination, actions that normally require an Environmental 

Assessment, and actions that normally require Environmental Impact Statements 

are codified in APHIS’ NEPA Implementing Procedures located in 7 CFR 372.5.

The three types of NEPA documentation are:

1. Categorical Exclusion

Categorical exclusions are classes of actions that do not have a significant

effect on the quality of the human environment and for which neither an 

environmental assessment (EA) nor an environmental impact statement 

(EIS) is required. Generally, the means through which adverse 

environmental impacts may be avoided or minimized have actually been 

built into the actions themselves (see 7 CFR 372.5(c)).

2. Environmental Assessment (EA)

An EA is a public document that succinctly presents information and 

analysis for the decision-maker of the proposed action. An EA can lead to 

the preparation of an environmental impact statement (EIS), a finding of 

no significant impact (FONSI), or the abandonment of a proposed action. 

3. Environmental Impact Statement (EIS)

In the event that a major Federal action may significantly affect the quality 

of the human environment (adverse or beneficial), or, the proposed action 

may result in public controversy, an EIS is prepared. 

Endangered Species Act (ESA) – This statute requires that programs consider 

their potential effects on federally protected species. The ESA requires programs 

to identify protected species and their habitat in or near program areas and 

documentation of how adverse effects to these species will be avoided. The 

documentation may require review and approval by the U.S. Fish and Wildlife 

Service and the National Marine Fisheries Service before program activities can 

begin. Knowingly violating this law can lead to criminal charges against 

individual staff members and program managers.

Migratory Bird Treaty Act – This statute requires that programs avoid harm to 
Last update 02FEB2022 Cydalima perspectalis 39

over 800 endemic bird species, eggs, and their nests. In some cases, permits may 

be available to capture birds, which require coordination with the U.S. Fish and 

Wildlife Service.

Clean Water Act – This statute requires various permits for work in wetlands 

and for potential discharges of program chemicals into water. This may require 

coordination with the Environmental Protection Agency, individual states, and the 

Army Corps of Engineers. Such permits would be required even if the pesticide 

label allows for direct application to water.

Tribal Consultation – This Executive Order requires formal government to 

government communication and interaction if a program might have substantial

direct effects on any federally-recognized Indian Nation. This process is often 

incorrectly included as part of the NEPA process, but must be completed prior to 

general public involvement under NEPA. Staff should be cognizant of the conflict 

that could arise when proposed federal actions intersect with tribal sovereignty. 

Tribal consultation is designed to identify and avoid such potential conflict.

National Historic Preservation Act – This statute requires programs to consider 

potential impacts on historic properties (such as buildings and archaeological 

sites) and requires coordination with local State Historic Preservation Offices.

Documentation under this act involves inventorying the project area for historic 

properties and determining what effects, if any, the project may have on historic 

properties. This process may require public involvement and comment prior to the 

start of program activities.

Coastal Zone Management Act – This statute requires coordination with states 

where programs may impact Coastal Zone Management Plans. Federal activities 

that may affect coastal resources are evaluated through a process called “federal 

consistency”. This process allows the public, local governments, Tribes, and state 

agencies an opportunity to review the federal action. The federal consistency 

process is administered individually by states with Coastal Zone Management 

Plans.

Environmental Justice – This Executive Order requires consideration of 

program impacts on minority and economically disadvantaged populations. 

Compliance is usually achieved within the NEPA documentation for a project. 

Programs are required to given consider if the actions might disproportionally 

impact minority or economically disadvantaged populations, and if so, how such 

impact will be avoided.

Protection of Children –This Executive Order requires federal agencies to 

identify, assess, and address environmental health risks and safety risks that may 

disproportionately affect children. If such a risk is identified, then measures must 
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be described and implemented to minimize such risks.
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Sampling

Insect Collection: Trapping

Pheromone traps

1. Trap set-up

♦ Use bucket traps (Unitraps) and follow the Bucket Trap Protocol.

♦ Use the pheromone combination 5:1 (Z)-11-hexadecenal:(E)-11-

hexadecenal.

♦ Load 0.000002 ounces of pheromone onto laminate pheromone dispensers 

(photo-setting resin on polypropylene film). Laminate dispensers are more 

effective than rubber septa dispensers.

♦ Change or re-charge dispensers with new pheromones every 1 to 2 weeks.

2. Trap placement and maintenance

♦ Traps should be placed at least 66 feet apart.

♦ Position traps roughly five feet above the soil surface.

♦ Place traps near hosts whenever possible, but away from foliage (See 

Bucket Trap Protocol).

♦ Service the trap every 2 weeks (Kim and Park, 2013).

Other available trapping techniques

♦ Light traps (Nacambo et al., 2014).

♦ Feeding attractant traps (phenylacetaldehyde–based and iso-amyl alcohol–

based lures) (Nagy et al., 2017).

Trapping in highly developed areas

Surveyors will need to consider the potential challenges when adjusting this 

survey to fit highly developed urban or residential landscapes with few suitable or 

accessible trapping sites. Optimizing the likelihood of capture by placing traps 

around possible hosts or in green areas near high-risk introduction points should 

remain the priority. High-risk areas will often include airports, warehouses, 

nurseries, or other sites that are likely to move high volumes of host plants and 

other products that can carry pests. Surveying around these high-risk areas will 

Appendix

B
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require on the ground assessments for access and site suitability as well as 

permissions from property owners to place and maintain traps. With permissions 

in place, trapping along roadsides, in parking lots, and in parks/green areas are 

likely to be essential for proper coverage in an urban landscape.

Sample Submission

Contact your State Plant Health Director for sample submission guidance.
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Plastic Bucket Trap 

Protocol

Plastic Bucket Trap 

Protocol

The plastic bucket trap is a long-lasting insect trap used in conjunction with a lure to monitor or 

detect various species of moths. The plastic bucket trap is the preferred trap for some moth species 

since it can catch large numbers of moths without damaging some of their identifying characters. 

The trap has four parts: 1) lid, 2) lure basket with cap, 3) funnel, and 4) bucket. The trap is available 

in various color combinations. For PPQ programs, the trap consists of a green lid, yellow funnel, and 

white bucket. Figure 1 is a photograph of a trap cut in half.

Figure 1 Plastic bucket trap cut in half to show its interior

Appendix

C
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Follow the steps below to prepare the bucket traps for use in the field.

1. Pheromone

The synthetic pheromone is embedded in a laminate (a small rubberized square as in Figure 2) or a 

septum (similar to a pencil eraser as in Figure 3) dispenser. Use gloves when handling lures and 

unwrap the lure from its packaging. When not in use, the lures should be stored in a freezer not used 

for food or drinks. Refer to the pheromone MSDS documents for storage and safety information. 

Figure 2 Laminate lure Figure 3 Septum lure

If you are using a laminate lure, you may attach the lure to a small paper clip and fold the clip so that 

the lure does not fall out of the basket (Figures 4 and 5). 

Figure 4 Laminate lure in Figure 5 Laminate lure in paper clip

paper clip inside lure basket
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If you are using a septum lure, place the septum inside the lure basket (Figure 6). Cover the basket 

with the lid and insert the basket through the circular opening on the center of the lid (Figure 7). If 

the cap no longer snaps snuggly into the trap lid opening, secure it with a piece of tape. 

Figure 6 Septum lure inside Figure 7 Lure basket with cap inserted through center of lid

lure basket

Note: To avoid cross-contamination from lure residue, do not reuse lure baskets for different 

species. Label the lure basket with the name of the moth species. 

2. Handle

Attach a wire handle to the lid through its two loops, as shown in Figures 8 and 9. A wire handle is 

usually included with each trap. If a handle is not included or needs to be replaced, make one with a 

12-inch long wire. You can also use string, but it does not last as long as the wire.

Figures 8 and 9 Wire handle attached to trap’s lid
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3. Trap Modifications to Improve Moth Specimen Quality

There are two trap modifications that may be used to improve moth specimen quality. In regions 

where little or no rain is expected, a sponge can be used to soak up small quantities of water (see 3a 

Sponge). This method does not require modifications to the trap. In regions where there is the 

possibility for significant rain, holes can be drilled into the bottom of the trap to allow for water to 

escape (see 3b Drain holes and wire screen). Surveyors are encouraged to use whichever method 

provides the highest quality moth specimens in their State’s unique climatic conditions. 

3a. Sponge

Place a dry cellulose sponge in the bottom of the trap, as shown in Figure 10. The sponge will 

absorb rainwater (except for extremely heavy amounts) that may enter the trap, keeping the moths 

somewhat dry. Sponges should be purchased locally. Purchase sponges without additives such as 

antibiotics, fragrances, etc.

Figure 10 Cellulose sponge (yellow) and insecticidal Figure 11 Bucket with five drilled holes

strip (red) inside the trap

3b. Drain holes and wire screen

Drain holes: Use a 3/16-inch drill bit to drill 4 to 5 holes in the bottom of the bucket to allow excess 

rainwater to drain (Figure 11). Holes should be drilled around the outer edge of the bucket to allow 

for proper drainage.

Wire mesh screen: Five-inch diameter wire mesh screens are available from the Survey Supply 

Ordering System in IPHIS (Product ID: 1381) (Figure 12). The wire mesh screen is placed in the 

bottom of the bucket trap to keep the moths from getting too wet from rainwater accumulated in the 

trap. When placed in the bucket trap, the screen is approximately 1-inch from the bottom of the 

bucket (Figure 13).
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Figure 12 5 inch wire mesh screen Figure 13 Wire mesh screen in bucket

Moving or handling the trap may cause the screen to tilt and allow specimens to fall below the screen. 

To improve the stability of the wire screen mesh use four #8 x 3⁄4 inch machine screws placed equally 

distance around the trap and 1-inch from the bottom of the plastic bucket (Tip: Use the small tabs on 

the inside of the bucket as a guide for placing screws 1-inch from bottom) (Figures 14 and 15). The 

use of screws allows for the easy removal of the wire mesh screen to collect specimens that may fall 

below (Figure 16).

Figure 14 Machine screws placed equal Figure 15 Machine screws shown one inch 

distance from each other around the bucket trap from the bottom of the bucket trap
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Figure 16 Wire mesh screen placed on 

machine screws inside bucket trap

4. Insecticidal Strips

Insecticidal strips (Figure 17) are placed in traps so that captured moths are killed quickly. This step 

is important for preserving specimen quality. The active ingredient in the strips is Dichlorvos, also 

known as DDVP and Vapona. The strip may be placed in the trap by: 1) attaching it to the side of the 

trap (as seen in Figure 10) or 2) placing it in the bottom of the trap or on top of the screen (if using 

the wire mesh screen modification). 

Figure 17 Insecticidal strip
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Note: The strip should be handled with gloves. Refer to the MSDS document for this product for 

safety information. Store unopened strips in a freezer that is not used for food or drink. 

The CAPS program has listed a conservative length of effectiveness of 8 weeks for the insecticidal 

strips. The manufacturers of insecticidal strips list an effectiveness of 12 weeks. Rain, wind, high 

heat, and large numbers of moths may reduce the potency of the insecticidal strips. Use the 8-week 

interval as a starting point, but you may change the interval (not to exceed 12 weeks) based on 

your state’s climate. If you find that moths are badly damaged (indicating they were not killed 

quickly), you should change the strips more frequently. In very hot and/or humid environments, it 

may be necessary to use two insecticidal strips, especially if traps are catching large quantities of 

moths.

5. Label the Trap

Attach a rain-proof printed label (Figure 18) or handwrite a note with a water-proof black marker on 

the bucket trap. It should indicate that the trap belongs to a state or a PPQ program. Include a phone 

number in case someone has concerns or questions about the trap. 

Figure 18 Label on the trap’s lid Figure 19 Trap set in open field 

and away from foliage

6. Placement of Traps

The traps function best when placed in the open, away from foliage, as illustrated in Figure 19. 

When hung under foliage, the 3-dimensional shape of the pheromone plume (chemical in the air) is 

disrupted and the effectiveness of the trap is reduced. Hang the traps from places such as greenhouse 

roofs or in the open using metal rods (see Fig. 19) or other materials.
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7. Trap Servicing

In the field, transfer the caught moths to labeled paper sample envelopes and store them in a cooler 

(Figures 20 and 21). Place them in a freezer overnight to kill any surviving specimens. Prior to 

screening, specimens can be stored in the freezer or in a cardboard box at room temperature. 

Note: Do not place samples in airtight containers, like plastic bags. The moths will rot, quickly 

degrading specimen quality and making identification difficult or impossible.

Figure 20 Moths placed in a paper envelope Figure 21 Moths stored in a cooler

8. Sample Submission

Prior to shipping, screen the samples. Remove any moths vastly different from the target moth and 

all other arthropods (beetles, flies, spiders). Write the approximate number of moths being submitted 

on PPQ Form 391. See Specimen Submission Guidance for Lepidoptera for instructions on 

preparing and submitting samples. 

Figure 22 Examples of small envelopes for shipping specimens
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Tip: The ideal envelopes for shipping samples are 3 1⁄2 x 6 1⁄2 inches and are commonly referred to as 

“coin envelopes” in office supply stores (Figure 22). Smaller or larger envelopes are acceptable as 

well. 

9. Trap Maintenance and Storage

The general recommendation for maintenance of the plastic bucket traps is to wash them 

occasionally with soap and water to keep them clean, and to store them indoors, or at least protected 

from sun, rain, and dust. Keep the wire handle and the wire screen in good condition. The traps can 

be used multiple times and for multiple species since the chemicals degrade quickly in outdoor 

conditions. These traps usually last more than five years.

Acknowledgments

This protocol has been designed to aid in the detection of exotic moths of concern by giving 

instructions on how to use generic plastic bucket traps. Photographs for figures 1-10, 17-19 and 21-

22 were taken by J. Brambila; trapping materials were supplied by R. Meagher. These instructions 

are primarily based on work by R. Meagher.

This aid was originally prepared by Julieta Brambila, Lisa Jackson (USDA APHIS PPQ), and Robert 

L. Meagher (USDA ARS CMAVE) in April 2010.

This aid was revised on October 10, 2014 by Julieta Brambila, Lisa Jackson, Douglas Restom 

Gaskill, Andrew Derksen (USDA APHIS PPQ), and Robert L. Meagher (USDA ARS SEA). 

Changes include:

• In the Pheromone section, added an image of the septum lure. Also added images on how to 

secure laminate lures into lure baskets with a paper clip.

• In the Pheromone section, added instructions on labeling lure baskets to avoid crosscontamination from lure residue. 

• In the Trap Modification section, noted that the sponge method may be easier and more effective 

than the screen method.

• In the Insecticidal strips, changed the recommended number of insecticidal strips to use in the 

trap from two to one.

• Changed the length of effectiveness of the insecticidal strips from 1 to 4 weeks to 8 weeks as a 

starting point; encourage states to evaluate what is effective in their climate.

• Added information on differing lengths of effectiveness of the insecticidal strips, depending on 

climate.

• In the Sample submission section, added two sample submission methods: sending moths in 

small envelopes or small boxes.

• In the Sample submission section, added images of the small manila envelopes.
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This aid was revised on April 30, 2021 by Heather Moylett, Todd Gilligan, and Christopher Pierce

(USDA APHIS PPQ); and reviewed by Mark Hollister (USDA APHIS PPQ). Changes include:

• In Trap Modifications to Improve Moth Specimen Quality section, expanded the drain holes 

and wire screen option to include how to stabilize the screen within the trap. Added figures 11-

16 to show modification. Photographs taken by Christopher Pierce.

• In Insecticidal Strips section: 

o Removed insecticidal strip placement option: “stapling it to a string and hung inside of 

the trap (if large trap catches are experienced).” The two remaining placement options are 

simpler and effective for even large sample volumes.

o Simplified “length of efficacy” guidance to improve clarity. 

• In Trap Servicing section, removed plastic bags as an option for sample storage and emphasized 

the use of paper envelopes for sample quality. Added figure 20. Photograph taken by Todd 

Gilligan. 

• In Sample Submission section, replaced written guidance with link to updated sample 

submission document maintained on the CAPS website. 
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Authors and Reviewers

Authors

Jennifer C. Cook, Ph.D., NCSU–CIPM

Thomas Culliney, USDA–APHIS–PPQ–PPRA

Colin F. Funaro, Ph.D., NCSU–CIPM

Jacob B. van Kretschmar, Ph.D., NCSU-CIPM

Editor

Rosemary Hallberg, NCSU–CIPM

Reviewers

Amanda Anderson, USDA–APHIS–PPQ–PPRA

Ignacio Baez, USDA–APHIS–PPQ–S&T

Audrey Bras, Ph.D., Swedish University of Agricultural Sciences

Steven Frank, Ph.D., North Carolina State University

Michelle L. Gray, USDA–APHIS–PPQ–PPRA

Wendy G. Marchant, Ph.D., USDA–APHIS–PPQ–PPRA

Godshen Pallipparambil, Ph.D., NCSU–CIPM

Olya Rysin, Ph.D., USDA–APHIS–PPQ–PPRA

Gregory S. Simmons, Ph.D., USDA–APHIS–PPQ–S&T

Michael J. Skvarla, Ph.D., Pennsylvania State University

Lindsey Thiessen, Ph.D., USDA–APHIS–PPQ–S&T

Ricardo Valdez, USDA–APHIS–PPQ–PER
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Contributors

Hui Fang, Ph.D., NCSU-CIPM

Global Pest and Disease Database Team