Document ID: EPA-HQ-OPP-2006-0657-0004
Agency: epa
Document Type: Supporting & Related Material
Title: 
Posted Date: 2006-10-11T04:00Z

SEQ CHAPTER \h \r 1 

UNITED STATES ENVIRONMENTAL PROTECTION AGENCY

WASHINGTON, D.C. 20460

	

		

		OFFICE OF PREVENTION,

PESTICIDES AND TOXIC 

									SUBSTANCES        

September 26, 2006

MEMORANDUM

SUBJECT:	Technical Review of Materials Supporting Arizona’s 24(c)
Applications [AZ05009 and AZ050010] to Replace Structured Non-Bt Cotton
Refuges with Sterile Insect Technology and 100% Bollgard and Bollgard II
Cotton (EPA Reg. No. 524-478 and 524-522) to meet their Special Need for
their Pink Bollworm Eradication Program [Submission dated August 3,
2006, MRID#s  469048-00 to -05]

TO:		Alan H. Reynolds, M.S., Entomologist 

		Regulatory Action Leader

		Microbial Pesticides Branch, Biopesticides and

		Pollution Prevention Division (7511P)

FROM:	Sharlene R. Matten, Ph.D., Biologist  /s/

		Technical Lead for Insect Resistance Management for Bt Crops 

		Microbial Pesticides Branch, Biopesticides and

		Pollution Prevention Division (7511P)

ACTION 

REQUESTED:	Provide a technical review of materials supporting
Arizona’s 24(c) Applications [AZ05009 and AZ050010] to replace the
required, structured, non-Bt cotton refuges with Sterile Insect
Technology and 100% Bollgard® and Bollgard II® cotton (EPA Reg. No.
524-478 and 524-522) to meet their Special Local Need for their Pink
Bollworm Eradication Program [submission dated August 3, 2006].

CONCLUSIONS AND RECOMMENDATIONS

The state of Arizona and the Arizona Cotton Research and Protection
Council submitted information to support continuation of the two Special
Local Need registrations (24(c) registrations [AZ-050009 and AZ-050010]
in place for the 2006 growing season.  A major concern with eliminating
the structured non-Bt cotton refuges and the possible use of 100% Bt
cotton is the possible increased risk of pink bollworm resistance to the
Bt proteins expressed in Bollgard and Bollgard II cotton varieties. 
This is the important issue for EPA.  Data from June 25 through July 22,
2006 were available for this submission due to the timing needed by EPA
to review the data and make a decision about the continuation of the
24(c) registrations.  

Based on the review of the preliminary pheromone trapping data, spatial
analysis, and simulation modeling, eliminating the structured non-Bt
cotton refuges, use of 100% Bt cotton, in combination with the release
of PBW sterile moths and pheromones with limited use of insecticides
during the PBW eradication program in Arizona, will not result in
increased risk of pink bollworm resistance to the Bt proteins expressed
in Bollgard and Bollgard II cotton varieties.  

The Kriging maps of native and sterile PBW populations in Arizona’s
eradication program from June 25 through July 22, 2006 are found in
Figures 6A-H (attached).  This analysis indicates that the sterile PBW
adult populations were more abundant, consistent and more widely
distributed than the native population.  The native populations were
limited to 1-5 moths per trap with 3-5 areas as “hot spots” (PBW
captures > 25) during this four week sampling period.  The sterile PBW
populations were more abundant with captures > 50 in many areas.  The
sterile populations were maintained at a steady level through daily
releases from aircraft during this program.  

Early results from the eradication program indicate that the sterile
releases have been quite successful in reducing native PBW populations. 
It is recommended that the spatial analysis be conducted on all of the
trapping data collected during the 2006 growing season and these results
be submitted to the Agency for review as a follow-up submission.  The
current analysis uses the centroid of the field to spatially locate the
pheromone trap for the Kriging analysis rather than the exact location
of the trap within the field.  It is also recommended that in future
years of the eradication program that the exact GPS coordinates of each
trap be provided for the spatial analysis to allow for greater precision
in the analysis.  This would allow a more precise examination of the
within field distribution of PBW.  For example, one would be able to
identify “hot spots” on one side of the field vs. another.  

The pheromone traps give a relative estimate of the population using
only male captures.  Other sampling methods, such as boll sampling, will
complement the pheromone trapping method to estimate PBW populations and
increase the precision of the spatial analysis.

Preliminary modeling using “pessimistic” (i.e., “worst-case”)
parameter assumptions predict that the four-year eradication program in
Arizona will suppress pink bollworm without creating a problem with
Cry1Ac resistance to Bt cotton.   The current simulations suggest that
the release of sterile moths in Bt cotton fields is important for
driving PBW population densities to extremely low levels.  However,
these are preliminary simulations and certain parameter values are best
estimates rather than actual field measurements.  Based on the summary
of actual sterile release rates (thru August 25, 2006; see Table 4), and
the actual percentage of Bt and non-Bt cotton fields see Table 1), the
range of input parameters used in the modeling simulations were accurate
(Table 8).  Additional modeling simulations should be done using actual
field values based on the data collected in 2006 and submitted to the
Agency as a supplemental submission.  

PBW resistance to the Cry2Ab2 toxin was not considered in either the
simulation modeling or DNA screening analyses.  Additional consideration
of PBW resistance to the Cry2Ab2 toxin would only be important if the
selection pressure dramatically increases in the next two years, i.e.,
much more Bollgard II planted in the eradication zone.  If some or all
of Arizona’s Bt cotton had two toxins, Cry1Ac + Cry2Ab, evolution of
resistance would be much less likely than it is with only Cry1Ac. 
Modeling resistance to cotton that produces only Cry1Ac is the most
pessimistic.  The modeling predictions (using only Cry1Ac resistance),
therefore, are conservative, i.e., they tend to overestimate resistance
risk.  Based on simulation models examining the likelihood of insect
resistance to pyramided toxins in Bt crops (e.g., Roush, 1998; Zhao et
al., 2005), even if Bollgard II acreage substantially increases, the
likelihood of PBW resistance to both the Cry1Ac and Cry2Ab2 toxins would
remain low during the four-year PBW eradication program in Arizona.  

It is recommended that the full season boll sampling and trapping data
(with the spatial analysis), DNA screening (molecular analysis) data,
and larval resistance monitoring data be submitted to the Agency for
review to confirm that there is a low likelihood of PBW resistance to
the Cry1Ac and Cry2Ab2 toxins expressed in Bollgard and Bollgard II,. 
Additional modeling simulations should be done with actual field data to
refine the model parameters as a way to partially field validate the
model and increase the accuracy of predictions that PBW resistance to Bt
cotton will be low during the planned PBW eradication program in
Arizona.  Follow-up resistance monitoring should also be done during the
eradication program and its aftermath.  

BACKGROUND AND SUMMARY OF THE SUBMISSION

The use of 100% Bollgard® and Bollgard II® cotton, in conjunction with
the use of pheromones, sterile insect technology, and limited
conventional insecticides for the purposes of pink bollworm
(Pectinophora gossypiella (Saunders), PBW) eradication, is a significant
change to the insect resistance management program.  The state of
Arizona (Department of Agriculture) and the Arizona Cotton Research and
Protection Council provided additional information in support of two
existing special local needs (SLN) registrations, AZ-050009 and
AZ-050010.  This information was submitted to address the conditions as
outlined in the Agency’s March 27, 2006 letter.  The specific data
needed were the result of the June 22, 2006 teleconference between
representatives of the EPA and representatives of the ACRPC, the state
of Arizona’s Department of Agriculture, and the National Cotton
Council.  As a result of this teleconference, the following data were
submitted to address the uncertainty of the effectiveness of the PBW
eradication program using sterile insect technology, 100% Bt cotton,
pheromones, and limited insecticide use.  

Data from GPS mapping of locations of all Bt cotton and all non-Bt
cotton plantings. The identity of individual grower fields is protected.
(Volume 1)

Data from systematic monitoring (weekly) of pink bollworm population in
eradication zones using pheromone traps and sampling of bolls were
provided. (Volume 2)

Data from systematic monitoring of resistance in moths collected in
pheromone traps within eradication zones (molecular analysis).  (Volume
3)

Data from the systematic resistance monitoring program for 2005/2006
larval populations.  (Volume 4)

Output from simulation modeling comparing population suppression vs.
resistance risk for the duration of the eradication program.  (Volume 5)

				

The Agency’s technical assessment of these data is the subject of this
review.

BI-NATIONAL PINK BOLLWORM ERADICATION PROGRAM BACKGROUND

The National Cotton Council estimates PBW costs western cotton producers
an estimated $21.6 million annually for prevention, control, and yield
losses.  An extensive review of PBW biology, ecology, and population
dynamics and integrated pest management options and approaches in the
southwestern United States is provided in Henneberry and Naranjo (1998).
 Pink bollworm eradication is possible in the southwestern United States
due to the following factors:

Limited hosts of PBW:  cotton and okra (non-preferred and extremely
limited in distribution) 

Availability of a very specific, highly efficient survey tool for
population detection and monitoring through trapping, i.e., Gossyplure
baited survey traps

Diverse and effective control mechanisms are available:  Bt cotton,
pheromone mating disruption, sterile insect technology, insecticides,
mandated plow down to lower over-wintering populations.

Economic feasibility.

Pink bollworm eradication efforts were begun in 2001 in the
Trans-Pecos/El Paso, TX area as part of a dual boll weevil and pink
bollworm eradication program.  A bi-national organization coordinates
the U.S. and Mexican bi-national pink bollworm eradication program. 
This program is a three-phase program.  Phase I was begun in 2001 in the
Trans-Pecos and El Paso areas of Texas with pheromone applications
followed by sterile moth releases in 2002.  Phase I was expanded to
include parts of New Mexico, and Chihuahua, Mexico in 2002 for a total
of approximately 160,000 acres of cotton. Phase II was initiated in 2006
and, in its entirety, will include western New Mexico and south-eastern
and central Arizona (240,000 acres).  Phase III is scheduled to begin in
2007 and will include western Arizona, southern California, and
northwestern Mexico (276,000 acres).  After five years, Phase I has
achieved a 99.4% reduction in both pink bollworm adults trap catches and
larval infestation percentages in the Trans-Pecos/El Paso area and
virtually a 99.9% reduction in both adult captures and larval
infestation percentages in South Central New Mexico and Chihuahua,
Mexico (USDA/APHIS presentation by Dr. Osama El-Lissy given at the
January 4-7, 2006 Beltwide Cotton Conferences, San Antonio, Texas). 
These results were achieved using Bollgard or Bollgard II cotton under
the current, structured refuge requirements.   

USDA-APHIS is responsible for technical support and coordination of the
bi-national program and for the administration of the sterile pink
bollworm moth component through a centralized management system.   The
program technologies are applied area-wide over a four-to-five (or more)
year period.   Local grower communities comprised of committees and/or
foundations are responsible for the daily implementation of program
operations, including mapping all cotton fields, tracking the
distribution of all transgenic Bt cotton fields, detection surveys, and
timely applications of pheromones on non-Bt cotton fields.

The San Joaquin Valley, California prevention program has 700,000 acres.
Sterile insect technology has been used for close to 40 years in the San
Joaquin Valley program and has successfully prevented the pink bollworm
from becoming established in that key cotton-producing area.  

Field data exist for the use of 100% Bollgard II in Hudspeth County,
Texas on approximately 2,100 acres in 2004.  The Agency approved this
amendment in April, 2004.  The Agency concluded in its technical review
of this amendment (Reynolds, 2004a) that the amendment was acceptable
and should not lead to pink bollworm resistance during the duration of
the eradication program because it was: 1) limited to less than 3,000
acres, 2) for one growing season, 3) the area is isolated, at least 30
miles, from any overwintering populations of tobacco budworm and cotton
bollworm, 4) Bollgard II expresses two high dose proteins (Cry1Ac and
Cry2Ab2) for pink bollworm control, thus reducing the likelihood of
resistance, 5) the eradication program included the release of sterile
pink bollworm males that will mate with any rare resistant survivors,
and finally, 6) intensive resistance monitoring was to be conducted in
this area.  The results of this field test were provided to the Agency
by the State of Arizona in December, 2005.  Based on the review of the
data, this field test was 100% effective based on the use of both bloom
and boll surveys (data collected in July and September, respectively by
Staten, Jenkins, and Walters, USDA/APHIS/PPQ).  Sterile pink bollworm
were released throughout the growing season after the first bloom at a
60: 1 sterile to native ratio to mate with an potential resistant
females emerging from Bollgard II cotton fields. There was no evidence
of a significant change in susceptibility to either Cry1Ac or Cry2Ab2
based on the resistance bioassays conducted by the University of
Arizona.  The Agency received no requests for any other amendments to
the Bollgard or Bollgard II cotton registrations for the use of sterile
moths and 100% Bollgard or Bollgard II cotton in 2005.  

Under the current terms and conditions for Bollgard and Bollgard II
cotton registrations (amended September 30, 2004), there are specific
structured refuge requirements to mitigate the likelihood of tobacco
budworm (Heliothis virescens), cotton bollworm (Helicoverpa zea), and
pink bollworm resistance to the Cry1Ac and Cry2Ab2 toxins:  1) 5%
external, unsprayed structured refuge (must be  within ½ mile of the Bt
fields, but ¼ mile or immediately adjacent is preferred), 2) 5%
embedded refuge (must be a least 150 feet wide, but preferably 300 feet
wide), 3) for pink bollworm, an in-field refuge strip refuge may be
used, with one row of non-Bt cotton planted for every six to ten rows of
Bt cotton, and 4) 20% external, sprayed structured refuge (must be
within one mile of the Bt fields, but ½ mile or closer is preferred).  
A community refuge option is also allowed for multiple growers within a
one-mile square area to use a combination of either the 5% external,
unsprayed refuge option and/or 20% external, sprayed refuge option.  The
embedded or in-field refuge options are not allowed under the community
refuge program.  Tobacco budworm and cotton bollworm are not insects of
concern in Arizona. 

SUMMARY OF ARIZONA’S SPECIAL LOCAL NEED REGISTRATIONS

The state of Arizona’s Special Local Need (FIFRA section 24(c))
Registrations (AZ-050009 and AZ-050010) permit the use of 100% Bollgard
and Bollgard II cotton varieties in a sanctioned pink bollworm
eradication program.  The eradication program (Phase 2) in Arizona was
initiated during the 2006 cotton growing season.  In May of 2004,
Arizona cotton growers passed by a 79% majority, a statewide referendum
that authorized the implementation of a pink bollworm eradication
program in Arizona.  The program is directed and executed by the Arizona
Cotton Research and Protection Council (ACRPC).  The program in Arizona
has been approved to run for a maximum of four years in each region
where it will be undertaken.  If eradication is not achieved after four
years, the effort will be discontinued.  If eradication is achieved,
some level of program surveillance and maintenance, comparable to that
currently in place for boll weevil, will be continued.  Support letters
for this program were provided by the University of Arizona scientists
involved in pink bollworm resistance management supporting the proposed
use of sterile insect technology refuges and 100% Bollgard and Bollgard
II cotton and by Monsanto who stated that they neither support nor
object to the program as presented.

In 2004, Arizona growers planted 73% Bollgard and Bollgard II cotton
varieties.  The ratio of adoption between Bollgard and Bollgard II
cotton varieties is approximately 95:5 based on review of Monsanto’s
annual sales information for the 2004 growing season (Matten, 2005). 
Bollgard II cotton varieties became available in the 2003 growing season
and are still fairly limited.  Arizona growers have experienced an
increase in returns of $5 million per year (± $2 million) as adopter
grains have outweighed non-adopter losses. Insecticide use in Arizona
cotton has declined since 1996 (the first year that Bollgard cotton was
available) from an average if six applications to an average of two
treatments in 2000.  Dramatic reductions in pink bollworm populations
have been shown to occur over large areas of Arizona in which Bt cotton
was high as noted in Carrière et al. (2003).  The state of Arizona,
therefore, believes that use of Bt cotton provides the potential to
eradicate pink bollworm during the four-years of the statutorily
approved program beginning with the 2006 growing season.

A major concern with eliminating the refuge is the possible development
of pink bollworm resistance to the Bt proteins expressed in Bollgard and
Bollgard II cotton varieties.  This is the important issue for EPA.  The
University of Arizona researchers have conducted statewide monitoring of
Bt resistance since 1997.  After nine years of use of Bt cotton, Bt
resistance alleles have been shown to be extremely rare in Arizona
(Tabashnik et al., 2006). In addition, the University of Arizona
researchers have shown that Cry1Ac resistance to Bt cotton is recessive.

The ACRPC will release sterile moths in all Bt and non-Bt fields within
active eradication zones.  These fields will be determined by ACRPC
personnel using GIS mapping techniques.  Cotton variety planting
information will be gathered from all growers and physically verified by
ACRPC supervisors as to the presence or absence of Bollgard and Bollgard
II cotton.  The ACRPC will maintain the theoretical 1:500 ratio of
susceptible (i.e., sterile moths) to potentially resistant PBW (this
ratio was recommended by the FIFRA SAP Panel in 1998) by releasing
sterile moths up to three times per week in Bt fields from first bloom
until defoliation in order to establish a rate of 20 sterile
moths/acre/day.  The sterile moths will be released directly over or
adjacent to the Bt field thus, putting them in the direct vicinity of
any developing resistant populations.

The ACRPC, in consultation with USDA and University experts, has devised
the following plan that replaces the biological function served by
non-Bt cotton refuges with artificially raised sterile pink bollworm
moths.

Planting of 100% Bollgard and Bollgard II cotton varieties will be
encouraged by eradication personnel, but not required.

Annual identification and GPS mapping of the locations of Bollgard and
Bollgard II  and non-Bt cotton plantings to provide precise information
with which to determine fields that do not satisfy the current refuge
requirements and to respond to future problems, should they arise. 

Non-Bollgard fields will be treated with a combination of pheromone and
insecticides.

When PBW populations in non-Bt fields will have declined sufficiently
for establishing suitable sterile/native moth ratios, sterile moths will
be released on non-Bollgard fields. Sterile moths may not be used in
non-Bt cotton until year two of the program in most areas of the state.

The State of Arizona proposes to release 20 sterile moths/acre three
times per week (49.4 sterile moths/hectare) in 100% Bt cotton fields to
satisfy the 500:1 susceptible/resistant moth ratio currently targeted by
the conventional refuge strategy.  The release in Bt cotton would start
at first bloom in year one in each zone of the eradication effort and
continue for the duration of the cotton growing season through the
length of the four-year program.  The sterile moth release rate over
non-Bt cotton fields is at a rate equivalent to 100 moths per acre per
day (three times per week) or higher (247 moths per hectare per day).

Systematic monitoring of resistance in moths detected in pheromone traps
within eradication zones, using pheromone traps and sampling of bolls,
will be conducted.

Systematic monitoring of resistance in moths detected in pheromone traps
within eradication zones, using molecular biological methods to detect
the mutations that have been shown to be associated with pink bollworm
resistance to Bt cotton in Arizona.

Statutory limitation of the duration of the eradication efforts to a
maximum of four years in each zone.

Monitoring of resistance using established methods based on collection,
culturing and bioassaying of larval populations of suitable density, if
and when they are found in any Bt cotton within eradication zones.

Continuation of a multi-agency pink bollworm group composed of the U.S.
Environmental Protection Agency, ACRPC, USDA, and University of Arizona
personnel, as well as grower and industry representatives, who will
closely monitor all scientific and regulatory aspects of the eradication
program and formulate case specific contingency plans for responding to
resistance development in eradication zones.

Primary emphasis on the use of environmentally benign pheromone methods
for control of pink bollworm in non-Bt fields within eradication zones
and secondary emphasis on limited use of conventional chemical
insecticides.

Lastly, an emergency response team that has been in place for nine years
to respond immediately to field developments will be used.  It is a
collaboration of ACRPC personnel under the consultation of University of
Arizona resistance monitoring experts.  Their task would be to verify,
document, and respond with remedial action to resistance problems.  The
plan for remedial action proposed by the Arizona multi-agency working
group and EPA is attached (Appendix 1).

EPA’s Review

EPA’s issues is whether eliminating the structured non-Bt cotton
refuges and the use of 100% Bt cotton will result in increased risk of
pink bollworm resistance to the Bt proteins expressed in Bollgard and
Bollgard II cotton varieties.  EPA’s review is divided into five
sections corresponding to the five volumes in the state of
Arizona’s/ACRPC’s submission.  The data were collected in the
certified PBW eradication zones for the 2006 season, June 25 through
July 22, 2006, except for the larval resistance monitoring data which
were based on 2005 collections.

Bt/non-Bt Mapping and Program Management Data

Geospatial mapping data of the location of all Bt cotton and all non-Bt
cotton plantings were collected to provide precise information as to the
location of all Bt and non-Bt fields.   Data were provided on the
management of these fields:  sterile release rates (per acre), program
applied pheromone treatment (per acre), program applied insecticide
treatments for PBW (per acre), pheromone trap captures expressed as
moths per trap per unit time by field, and boll infestation levels
expressed as larvae per 100 bolls in fields selected for program
evaluation.  The ACRPC supplied four maps illustrating the Bt/non-Bt
field locations in Central and Eastern Arizona (Maricopa, Pinal, Pima,
Graham, Cochise, and Greenlee counties) and the sterile moth release
zone (Figures 1-4 from Volume 1 of the submission, MRID# 469048-01)
listed below.  

Map 1A (Figure 1).  Bt/non-Bt field locations - Central Arizona

Map 1B (Figure 2).  Bt/non-Bt field locations - Eastern Arizona

Map 2 (Figure 3A and B).  Expandable format of Map 1A (Fig. 1) and 1B
(Fig. 2) (with individual fields numbered to match trapping/treatment
data bases

Map 3 (Figure 4).  Sterile moth release zones with expanded inset maps
including flight path data.

The total acreage represented by Bt fields and non-Bt fields in the 2006
PBW eradication program in each of the six counties in Arizona is shown
in Table 1.  There were a total of 165,632.95 acres in the 2006 PBW
eradication program in Arizona.  Ninety-two percent of the total cotton
acres in the program were in Pinal, Maricopa, and Graham counties with
sixty percent of the total cotton acres in Pinal county.

Table 1.  2006 Bt and non-Bt Cotton Acreage Per County in Arizona Pink
Bollworm Eradication Program [Table from Letter L. Antilla, ACRPC to S.
Matten, USEPA/OPP/BPPD, dated September 14, 2006]

 

There were 4,626 total fields in the eradication zone:  334 non-Bt
fields (6.92%) and 4,292 Bt fields (93.08%) (Table 2).  Each field is
numbered.  These fields are the target areas for the sterile moth
releases, pheromone, and insecticide treatments. There were a total of
4,541 pheromone traps placed in all fields with 3,541 pheromone traps
placed in Bt fields and 1,000 pheromone traps place in non-Bt fields. 
The number of traps per field ranged from 0 to 14.  The scheme for using
the trapping and map data is shown in Figure 5.   

Table 2.  2006 Bt  and non-Bt  Cotton Total Acres, Percent, Total
Fields, and Total Traps in Arizona Pink Bollworm Eradication Program
[Modified Table from Letter L. Antilla, ACRPC to S. Matten,
USEPA/OPP/BPPD, dated September 14, 2006]

 

The program applied pheromone and insecticide treatment data (June 25
through July 22, 2006) are summarized in Table 3.  A total of 806.8
total acres (10 fields, some more than once) were treated.  This
represents only a small fraction, 0.2%, of the total cotton fields in
the program.  The PBW pheromone rope (PB-Rope L®
((Z,Z)-7,11-Hexadecadien-1-yl Acetate 46.7% (Z,E)-7,11-Hexadecadien-1-yl
Acetate 44.1%), Gossyplure, Pheromone) was applied to all non-Bt cotton
at the sixth to seventh true leaf stage at a rate of 200 dispensers per
acre (approx. 11,500 acres were treated).   The dispensers are applied
several different ways.  Dispensers are either hand tied to the plants,
wrapped on a bamboo stick and placed in the planted row by hand or,
wrapped on a bamboo stick and mechanically inserted into the planted
row. PB-Rope L is a 60-90 pheromone mating disruption treatment.  The
following other insecticides were used: Dual® and Lock-On®.  Dual
indicates a dual treatment including a mating disruption pheromone
constituent such as NoMate PBW fiber ((Z,Z)-7,11 -Hexadecadien-1-01
acetate 3 .80% + (Z,E)-7,1 1- Hexadecadien-1-01 acetate 3.80%)
(Gossyplure, Pheromone) or NoMate PBW MEC ((Z,Z)-7,11- Hexadecadien-1-
yl Acetate 10.0% + (Z,E)-7,11- Hexadecadien-1 -yl Acetate 10.0%)
(Gossyplure, Pheromone) (Microencapsulated concentrate) and a chemical
component such as Lock-On.  Lock-On is a formulation of
microencapsulated chlorpyrifos.

Table 3.  ARIZONA COTTON RESEARCH AND PROTECTION COUNCIL

	PROGRAM APPLIED PHEROMONE AND INSECTICIDE 

	TREATMENT DATA

	 JUNE 25 THROUGH JULY 22, 2006

	Field#	Type	Pheremone	Insecticide	Acres Sprayed

	2367	DUAL	MEC	LOCK-ON	41

	2367	DUAL	MEC	LOCK-ON	146.5

	2367	DUAL	MEC	LOCK-ON	146.5

	2366	DUAL	MEC	LOCK-ON	27.1

	2366	DUAL	MEC	LOCK-ON	27.1

	2365	DUAL	MEC	LOCK-ON	27.9

	2365	DUAL	MEC	LOCK-ON	27.9

	2364	DUAL	MEC	LOCK-ON	28.3

	2364	DUAL	MEC	LOCK-ON	28.3

	2363	DUAL	MEC	LOCK-ON	28.2

	2363	DUAL	MEC	LOCK-ON	28.2

	2362	DUAL	MEC	LOCK-ON	28.1

	2362	DUAL	MEC	LOCK-ON	28.1

	2359	DUAL	MEC	LOCK-ON	39.9

	2359	DUAL	MEC	LOCK-ON	39.9

	2358	DUAL	MEC	LOCK-ON	33

	2358	DUAL	MEC	LOCK-ON	33

	2187	DUAL	MEC	LOCK-ON	24

	2186	DUAL	MEC	LOCK-ON	23.8

	GRAND TOTAL ACRES SPRAYED	806.8

	Pheromone rope or "PBROPE" was applied to all NON-BT cotton at the
sixth to 

	seventh true leaf stage. The rope was applied at the rate of two
hundred dispensers 

	per acre. 11,537.96 acres were treated with PBROPE from May 7, 2006 to
July 10, 

	2006.

The success of using sterile moths in the PBW eradication program is
dependent on whether the sterile moths are as competitive as the native
moths.  Miller et al. (1994) in their research found that both male and
female sterile PBW moths are comparable to native moths in their mating
responses.  Studies by Tabashnik et al. (1999) indicate that both native
and sterile male PBW can move up to 400 meters (approx. ¼ mile) from
non-Bt and Bt cotton.  This means that sterile releases within 400
meters of Bt cotton field should provide a sufficient PBW populations of
both males and females within the boundary for effective interaction
with native moths whether they are Bt-susceptible or resistant.  Further
research conducted by the California Department of Food and Agriculture
(Keaveny et al., 2006 in Vol. 2, MRID#  469048-02) indicates that
sterile moths release in one mile (approx. 1600 meters) corridors move
effectively at least one mile offsite for potential encounters with
native moths.  The movement information is important for use in the
Kriging analysis and for determining the sterile release flight
corridors.  

The protocol for the sterile moth releases is as follows:

Non-Bt Cotton:  Sterile moth (sterile insect technology, SIT) releases
are made three times per week directly over non-Bt fields at a rate
equivalent to 100 moths per acre per day or higher (247 moths per
hectare per day or higher).

Bt Cotton:  SIT releases are made 2-3 times per week along one mile
corridors over Bt fields at a rate equivalent to 20 moths per acre per
day or higher (49.4 moths per hectare per day or higher).  This release
rate represents a minimum of two times the USDA/APHIS release rate for
Bollgard and Bollgard II referenced in Arizona’s 24(c) labels. 
Corridors are offset by one half mile (approx. 800 meters) on alternate
release days to ensure that sterile moth populations are maintained
within one quarter mile (approx. 400 meters) of Bt fields at all times
throughout the season.

All SIT releases are monitored through GPS assisted guidance systems
which produce flight recordings which are downloaded, printed and
reviewed daily.  These include flight paths and color coded designations
of all release operations.

Arizona:  Allocation of sterile moths is 70 million moths per week for
an overall average of 10 million moths per day.  Releases are made seven
days a week and cover the period of May 1st through October 15, 2006.

Pheromone traps on all Bt and non-Bt fields are serviced weekly and
counts of native vs. sterile moths are recorded by trained ID personnel
at each field office.  Because sterile moths are reared on a diet
containing red dye, either visual or simple chemical assays separate
sterile moths from native moths.  



Information on Actual Sterile Release Rates (personal communication from
B.Tabashnik, U of Arizona to S. Matten, USEPA/OPP/BPPD, dated September
14, 2006)

Table 4.  Sterile Release Rates through 8/25/06

Non-Bt cotton, 3 releases per week (1 release per 2.3 days) 

Mean to date = 251 moths per acre per release (621 per ha per release) 

251 moths per acre per release X 3 releases per week = 753 moths per
acre per week 

753 moths per acre per week divided by 7 days per week = 108 moths per
acre per day 

Bt cotton, 3 releases per week (1 release per 3 days) 

Mean to date = 53.1 moths per acre per release (131 per ha per release) 

53.1 moths per acre per release X 3 releases per week = 159.3 moths per
acre per week 

159.3 moths per acre per week divided by 7 days per week = 22.8 moths
per acre per day 

If non-Bt = 7% of acreage, Bt =  93% of acreage, 108 X 0.07 + 22.8 X
0.93 = 7.56 + 21.2 = 

actual mean release rate = 28.8 moths per acre (71.1 per ha) per day 

Note:  Production of 70 million moths per week /165,000 acres of cotton
= 420 moths per acre per week = potential mean release rate = 60 moths
per acre per day (estimated mean is half of this) 

Discussion

The Bt and non-Bt cotton field maps and the sterile moth release
protocol are “acceptable.”  Actual sterile release rates in Bt and
non-Bt fields are as expected (see Table 4 above).  

Pink Bollworm Trapping and Boll Sampling Data

Data from systematic weekly monitoring of PBW populations in eradication
zones using pheromone traps and sampling of bolls was provided for the
period June 25-July 22, 2006.  Conclusions based on these data will be
preliminary given the short period in which the trapping and boll
sampling data represented.  Weekly monitoring will be continued through
October 15, 2006.  Cumulative trapping, treatment and sterile release
data should be provided to the Agency at the end of the 2006 cotton
growing season as a follow-up submission.

Adult PBW populations were monitored using a program-wide standard PBW
delta trap baited with PBW sex (male) pheromone (Gossyplure, Pheromone,
Hexadecadienyl acetate).  Traps were placed in each cotton (Bt and
non-Bt) in the eradication zone (six Arizona counties noted above).  
The number of traps per field varied based on the size of the field, and
the type of cotton grown.  The number of traps ranged from 0 to 14.  A
field may have no trap located in it because there was one central trap
that covered several small fields.  The average number of moths per trap
was calculated for each cotton field by dividing the total capture
(steriles or natives) by the number of traps.   A total of 4,541 traps
were placed in 4,626 cotton fields in 2006 (see Table 2 above).  Traps
were checked weekly by program personnel and numbers of male moths
(native and sterile) were recorded.  A detailed description of the
trapping and sampling methodology is found in Appendix 2.  

  

Raw data listing weekly program trapping for all fields, the number of
traps per field and the total number of moths counted each week, for the
period of June 25 through July 22, 2006 are provided in Volume 2, Table
1 (146 pages) of the submission (MRID# 469048-02).  Some trapping dates
have missing information (i.e., null data).  No information as to why
trap counts are missing for these dates is provided.  The average number
of native and sterile moths per field for the four week combined date
range of June 25 to July 22, 2006 are provided in Volume 2, Table 2 (100
pages) of the submission (MRID# 469048-02).  The weekly PBW eradication
program averages by cotton type during the period of June 25 to July 22,
2006 are provided below in Table 5.

Table 5.  Weekly PBW (Natives and Steriles) Eradication Program Averages
by Cotton Type 6/25-7/22/2006. (Dates correspond to the start date of
the trapping week)

Cotton Type	Nat/Trp

6-25	Nat/Trp

7-02	Nat/Trp

7-09	Nat/Trp 7-16	Str/Trp

6-25	Str/Trp

7-02	Str/Trp

7-09	Str/Trp

7-16

Bt	3.20	1.92	2.75	1.83	20.50	29.77	46.48	29.94

Non-Bt	0.18	0.05	0.11	0.10	0.73	1.09	1.53	1.16

The average weekly sterile to native moth ratio for the Bt fields for
each trapping week varied from 6.4 to 16.9.  The average weekly sterile
to native moth ratio for the non-Bt fields for each trapping week varied
from 4.1 to 21.8.  Captures (steriles and natives) were always greater
in Bt fields than non-Bt fields.  

Spatial Analysis

The spatial analysis of the trapping data was conducted by David
Bartels, USDA-APHIS-PPQ-CPHST (Edinburg, TX) and Michelle Walters,
USDA-APHIS-PPQ-CPHST (Phoenix, AZ) (see details of the method described
in Bartels and Walters, 2006 in Volume 2 of the submission, MRID#
469048-02).  To present the trapping data as a predicted surface of PBW
numbers (i.e., trap counts at a particular point), the Kriging method
was used to calculate a predicted value for areas between the known
values of each field.  Kriging is a validated geostatistical method used
to estimate the optimal interpolation of these points across the spatial
domain.  This method handles spatial autocorrelation and is not
sensitive to uneven sampling in specific areas, such as the distribution
of cotton fields in the eradication program.  Ordinary Kriging using a
spherical model was applied to trap counts for each week (see Volume 2,
Table 1 of the submission, MRID# 469048-02) to develop a predictive
surface model encompassing the cotton fields.  Kriging constructs a
weighted moving average that estimates the value of a spatially
distributed variable from adjacent values while considering the
interdependence.  Kriging results in a smoothing effect in which high
original values are underestimated and low original values are
overestimated.  It is a best linear unbiased estimator because it
minimizes the variance of the estimation errors.    

To create a point for the trap captures, the center of each cotton field
containing a trap was calculated using its geographic boundary.  Krigid
surfaces were generated from a total of 3,472 center points from the
cotton fields (used weekly trapping data from June 25 to July 22, 2006,
Vol. 2, Table 1, MRID#  469048-02).  Ideally, one would use the exact
GPS coordinates for the specific traps, but this information was not
available for the preliminary spatial analysis.  A two kilometer range
of influence from the center point of the field was used so that each
field’s data is only affected by other fields within approximately one
kilometer of the outside border.  This one kilometer limit reflected the
perceived day to day movement of PBW adults in Bt cotton fields and
limits the mathematical influence of a “hot” field on a large area. 
Trap values were truncated to 100 moths/field/week as this indicates a
“hot” field biologically and also, because a weighted average is
used in Kriging, capping the high value limits the undue graphical
influence of a single field.  Traps with greater than 100 PBW moths are
potentially more unreliable because the efficiency of the trap declines
once it fills up with moths.  

 Results and Discussion

The Kriging maps of native and sterile PBW populations in Arizona’s
eradication program from June 25 through July 22, 2006 are found in
Figures 6A-H (attached).  This analysis indicates that the sterile PBW
adult populations were more abundant, consistent and more widely
distributed than the native population.  The native populations were
limited to 1-5 moths per trap with 3-5 areas as “hot spots” (PBW
captures > 25) during this four week sampling period.  The sterile PBW
populations were more abundant with captures > 50 in many areas.  The
sterile populations were maintained at a steady level through daily
releases from aircraft during this program.  

Early results from the eradication program indicate that the sterile
releases have been quite successful in reducing native PBW populations. 
It is recommended that the spatial analysis be conducted on all of the
trapping data collected during the 2006 growing season and these results
be submitted to the Agency for review.  The current analysis uses the
centroid of the field to spatially locate the pheromone trap for the
Kriging analysis rather than the exact location of the trap within the
field.  It is also recommended that in future years of the eradication
program that the exact GPS coordinates of each trap be provided for the
spatial analysis to allow for greater precision in the analysis.  This
would allow a more precise examination of the within field distribution
of PBW.  For example, one would be able to identify “hot spots” on
one side of the field vs. another.  

The pheromone traps give a relative estimate of the population using
only male captures.  Other sampling methods, such as boll sampling, will
complement the pheromone trapping method to estimate PBW populations and
increase the precision of the spatial analysis.

Boll Sampling

Details of the boll sampling methodology are found in Appendix 2. 
Preliminary boll sampling data from July 16 to July 22, 2006 were
provided to the Agency (see Volume 2, Table 4 in MRID# 469048-02).  A
total of 43 fields were selected at random (23 non-Bt/20 Bt).  Attempts
were made wherever possible to select Bt/non-Bt pairs that were a mile
apart or less.  A total of 100 bolls from each of four field quadrants
comprised the sample.  These were then examined under magnification for
signs of PBW. Only 26 fields were able to be sampled at this early date.
 Only 1 boll from one non-Bt field was found to be infested with 1
larva.  The total infestation rate for all bolls analyzed was 0.04%. 

Discussion

Boll sampling this early in the season is preliminary.  It is
recommended that the boll sampling data collected during the entire 2006
growing season be provided to the Agency for review as a supplemental
submission.

Resistance Monitoring Data from Moth Collected in Pheromone Traps
(Molecular Analysis)

It has been shown in previous analyses that in laboratory-selected
strains of pink bollworm and at least two other major lepidopteran pests
of cotton, mutations in a cadherin gene are tightly linked with
recessive resistance to Cry1Ac (Gahan et al. 2001, Morin et al. 2003, Xu
et al. 2005). Previous work conducted at the University of Arizona has
identified three mutant alleles (r1, r2, and r3) of a cadherin gene
(BtR) are associated with resistance to Cry1Ac and survival on Bt cotton
(Morin et al., 2003, 2004; Tabashnik et al. 2004, 2005b).  Each r allele
has a deletion predicted to eliminate at least eight amino acids
upstream of the putative Cry1Ac-binding region of the cadherin protein
(Morin et al., 2003).  A PCR-method was developed to detect the r1, r2,
and r3 alleles in PBW (Morin et al., 2004).  This PCR-method was used to
screen for the three r alleles in PBW sampled from 59 cotton fields in
Arizona, California, and Texas during 2001-2005 (Tabashnik et al.,
2006).  No r alleles were detected in 5,571 field-derived insects.  

Methods for detecting false negatives, false positives, and
non-detection are described in detail in Tabashnik et al. (2006).  These
methods are also detailed in Appendix 3.     

In brief, false negatives are possible from three causes: i) The PCR
reaction is not working properly, ii) The cadherin DNA of field-sampled
insects is not amplified, iii) The PCR is working and cadherin DNA is
amplified from field samples, yet r alleles are present and are not
detected.  

To detect false positives, all tests of field samples include blanks,
which are gel lanes containing all of the PCR reagents, but no DNA.  If
a blank yields a positive result, this indicates contamination (i.e., a
false positive).  In this case, PCR reaction conditions are corrected
and the field samples are retested.  Results are included in the data
analysis only if the blanks do not yield positive results.  

Of the 5,571 field-sampled insects tested in Tabashnik et al. (2006),
none yielded positive results.  Thus, the problem of false positives is
minimal to nil.  When a pool of field-sampled insects yields a positive
result for an r allele (e.g., r2), each individual in the pool will be
tested separately to verify the positive result and to more precisely
estimate the frequency of resistance in the pool.

As described in Tabashnik et al. (2006), the likelihood of non-detection
is estimated as follows:

“The probability of detecting no r alleles in a sample of N
individuals was calculated as (1-[F X D])2N X A, where F is the
frequency of resistance alleles, D is the probability of detecting an r
allele present in screened individuals (0.97, based on the data from
blind controls), 2N is the number of alleles screened, and A is the
probability of amplifiable cadherin DNA occurring in field-sample
insects (estimated as 0.986, based on the proportion of positive results
for amplification of a conserved sequence in 835 insects tested
individually).  We assumed that the probability of an r allele occurring
was an independent event at each cadherin allele screened.  For example,
with an r allele frequency of 0.001, the probability of detecting no r
alleles in the sample of 5,571 individuals (11,142 alleles) is 0.000023
= (1- [0.001 X 0.97])11,142 X 0.986.  Analogously, with an r allele
frequency of 0.0003, the probability of detecting no r alleles in the
sample of 5,571 individuals is 0.041 = (1 – [0.0003 X 0.97])11,142 X
0.986.”

Results

As stated above, a series of positive and negative controls were used to
make sure the screening method was performing as expected.  Testing was
on insects in pools of 11 samples or fewer to minimize the chances of
missing r alleles.  Ninety-seven percent of r alleles were detected in
the blind positive control samples in these fields.  Statistical
analysis was used to calculate the probability of detecting no r alleles
in a sample of N individuals.  Based on the 98.6% amplification of
cadherin DNA and a detection rate of 97% for r alleles, the estimated
probability is <0.0001 that the frequency of r alleles in the field was
equal to or greater than 0.001 (Tabashnik et al, 2006).  The estimated
probability is <0.05 that the frequency of r alleles in the field was
equal to or greater than 0.0003.  Using this estimate and assuming
Hardy-Weinberg equilibrium, the estimated frequency of rr is less than 1
in 10 million (0.0003 X 0.0003).  Results from these analyses indicate
that r alleles for Cry1Ac resistance are rare.  

The resistance allele frequency estimated from bioassays conducted on
collections from 2001 to 2005 is somewhat higher than the estimate based
on DNA screening during the same time period.  The bioassay data for
2001 to 2004 were summarized in Tabashnik et al. (2005a) and have been
reviewed by the Agency (Reynolds, 2004b, c, 2005, 2006).  The mean
yearly estimate resistance allele frequency from 2001-2004 bioassays is
0.024 (range = 0 to 0.075, 95% confidence interval = 0 to 0.062). 
Tabashnik et al. (2006) suggest that the difference in the estimates
between bioassays and DNA screening in some years might be due to an
overestimation of the estimates using bioassays and underestimation of
the estimates by DNA screening, or possibly both.  It should be noted,
however, that there may be other mutant alleles of the cadherin gene or
other genes associated with resistance to Cry1Ac and survival on Bt
cotton that have not yet been identified. 

Plans for DNA Screening of Pink Bollworm in Arizona 2006

A DNA screening plan was developed by Dr. Tim Dennehy and Dr. Bruce
Tabashnik, University of Arizona to screen 500 field-sampled insects (50
or more per site from 10 sites) in Arizona for the three known cadherin
resistance alleles (r1, r2, and r3) using the PCR method described by
Tabashnik et al. (2006).  A brief description was found in Volume 3,
p.1, MRID# 469048-03).  Details of this screening plan are provided
below (personal communication, T. Dennehy, U. of Arizona to S. Matten,
USEPA/OPP/BPPD, dated September 14, 2006).  Low numbers may limit the
number of insects that can be collected and screened from the
eradication zone.  If possible, at least 300 field-sampled insects from
at least 6 sites in the eradication zone will be screened.  Special
effort will be made to collect insects for DNA screening from any areas
in which trapping data show unexpectedly high numbers of native moths in
the eradication zone.  Large numbers of native moths (50-100) needed for
these analyses are not normally available until September or October
when late season moth flights occur. 

Sampling methods:  As described by Tabashnik et al. (2006), insects for
DNA screening will be sampled from bolls and from traps baited with sex
pheromone:

“Cotton bolls were sampled from 19 cotton fields (18 in Arizona and 1
in California) from 2001 to 2005 as described by Dennehy et al. (2004). 
At each site, 300 to 2,000 bolls were collected from non-Bt cotton
fields near Bt cotton fields. Bolls were taken to the University of
Arizona Extension Arthropod Resistance Management Laboratory in Tucson. 
We obtained pink bollworm by collecting fourth instars that exited bolls
and by opening bolls and removing larvae found inside.” 

“Pink bollworm males were collected in sticky traps baited with female
sex pheromone (Tabashnik et al. 1999) in 40 cotton fields (36 in
Arizona, 3 in California, and 1 in Texas) from 2003 to 2005.  At each
site, several traps were placed around the perimeter of a cotton field,
collected after 1 to 2 days, and brought to the laboratory.  Live males
that showed normal movement of appendages were removed from traps using
wooden toothpicks.  A new toothpick was used for each male to avoid
cross-contamination.”

Progress and sampling plans:  

1.  In 2006, ACRPC proposed to identify up to five sample areas in which
elevated native Bt fields may suggest a resistance threat.  As of July
22, 2006, only one group of fields gave any indication of increased
native moth levels (field #s 4335-4338).   Approximately 80 male moths
were caught.  These moths will be screened using the three DNA markers
and PCR.

2. To produce strains for bioassay testing, bolls have been sampled from
four sites (two in the eradication zone, two outside the eradication
zone).  If numbers are sufficient, subsamples of 50 insects per site
collected directly from bolls will be screened for r alleles.

3. Trapping at 10 sites in the eradication zone will be done to obtain
males for PCR screening in late September.  If this sampling does not
yield enough males, trapping will be repeated in mid- to late October.

Discussion

DNA screening analyses of insects sampled from the field in 2006 could
not be performed prior to the August submission required by EPA to
support the two 24(c) registrations.  No information can yet be gained
about the presence or absence of the three mutations associated with
Cry1Ac resistance and survival on Bt cotton until the molecular analyses
are performed later this year. However, based on previous DNA screening
analyses of insects sampled from 2001 to 2005 (Tabashnik et al., 2006),
no individuals were identified as having these specific r allelic
mutations for Cry1Ac resistance.   It is recommended that the results
of these analyses be provided to the Agency. The methods for conducting
the molecular analyses for detecting the three mutations associated with
Cry1Ac resistance and survival on Bt cotton are “acceptable.”  If
alleles other than cadherin mutants r1, r2, and r3 confer pink bollworm
resistance to Bt cotton, the results using these three DNA markers DNA
could underestimate the frequency of resistance.    However, Tabashnik
et al. (2006) concluded that additional resistance alleles at the
cadherin locus or other loci are rarer than the three known resistance
alleles because such additional alleles have not been discovered in
extensive testing of several laboratory-selected resistant strains.  
DNA screening based solely on males caught in pheromone traps could
underestimate resistance allele frequency if the probability of capture
in traps was lower for rr or rs males than for ss males.  However,
experiments conducted in large cages (64 m3) in the field refuted this
argument (Carrière et al., 2006).  Furthermore, DNA screening of pink
bollworm of both sexes from bolls also detected no r alleles.  It is
recommended that the results of the 2006 DNA screening be submitted to
the Agency for review as a supplemental submission. 

No information is available about the nature of potential PBW resistance
to the Cry2Ab2 toxin.  Assuming adoption of Bollgard II will continue
to increase; understanding the genetics and possible mechanisms of
resistance to the Cry2Ab2 toxin will become more important.  It is
recommended that the genetics and potential mechanisms of PBW resistance
to the Cry2Ab2 toxin be studied.  Specific DNA markers would need to be
developed based on PBW resistance to Cry2Ab2.  These specific markers
would then be used in the DNA screening program.  One caveat is that the
adoption of Bollgard II has been low in Arizona (i.e., 95:5 ratio of
Bollgard:Bollgard II adoption) and the PBW eradication program is
limited to four-years so the need for such information may not be
crucial unless the adoption of Bollgard II, and consequently, Cry2Ab2
selection pressure, increases dramatically in the next couple of years. 
 However, as noted in the modeling section below, PBW resistance to both
Cry2Ab2 and Cry1Ac would be unlikely during the four-year eradication
program.  No information on the adoption of the relative adoption of
Bollgard and Bollgard II in the eradication zone for the 2006 growing
season is yet available.

  

Resistance Monitoring Data for 2005/2006 Larval Population

Annual Bt cotton resistance monitoring data for pink bollworm is a
requirement of the Bollgard and Bollgard II registrations.  All of the
Cry1Ac and Cry2Ab2 resistance monitoring data for pink bollworm larval
populations through the 2004 growing season have been previously
reviewed by the Agency (see EPA 2001; Reynolds 2004b (review of
2001/2002 growing season collections), 2005 (review of 2003 growing
season collections), 2006 (review of 2004 growing season collections)). 
  In these reviews, it was concluded that through the 2004 season, there
was no evidence of pink bollworm resistance to the Cry1Ac or Cry2Ab2
delta-endotoxins produced by Bollgard or Bollgard II cotton cultivars
under field situations.  The state of Arizona submitted a preliminary
monitoring report of the bioassays for the 2005 collections of pink
bollworm (Dennehy et al., 2006; in Vol. 4, MRID# 469048-04).  These data
are discussed below.

The 2005 monitoring work for pink bollworm was conducted in Arizona by
researchers at the University of Arizona and the ACRPC, who have been
conducting the work since 1997 (see Volume 4, MRID# 469048-04).  The
methodology for the 2005 PBW assays was largely the same as in previous
years and utilized artificial diet tests with a 21-day observation
period.  A discriminating dose type approach was used in which PBW
mortality was assessed to two test concentrations for both Cry1Ac and
Cry2Ab2.  Baseline susceptibility (i.e. a LC50 or similar measure) was
not determined.  The two test concentrations of Cry1Ac and Cry2Ab2 used
were 1.0 µg/ml and 10 µg/ml.  Negative controls (no toxin) were also
tested.  The Cry1Ac toxin used in the assays was obtained from Dow
AgroSciences (MVP-II Bioinsecticide) while the Cry2Ab2 toxin was
obtained from freeze dried corn powder provided by Monsanto.  These
toxin sources were also used for the 2005 monitoring.   In addition to
the laboratory bioassays, field efficacy was assessed in 2005 using
adjacent pairs of Bt and non-Bt fields at 44 Arizona locations. 

	2005 Sampling and Assays

PBW were collected as larvae (from bolls brought to the laboratory) from
Arizona (12 sites) and California (4 sites).  No samples were collected
in New Mexico and Texas due to ongoing PBW eradication efforts in those
areas.  At each location, 300 to 2,000 bolls were collected from non-Bt
cotton fields.  Laboratory cultures were established with ≥100 PBW
from each collection site.   A susceptible laboratory strain was also
used as an internal standard for the experiments.  Additionally, for the
Cry2Ab2 tests, a Cry1Ac-resistant PBW laboratory colony was included. 
Fourth instar larvae emerging from bolls were reared to adulthood to
produce progeny for testing (F2 - F8 progeny were used in the tests). 
The bioassays were conducted with artificial diet incorporated with the
two test concentrations (an untreated control was also used).  Neonate
larvae were placed in one ounce cups with diet and observed for 21 days.
 Larvae that failed to develop past the third instar by the end of the
test were considered “dead” and Abbott’s formula was used to
obtain corrected mortality scores (i.e. to justify mortality in the
control groups).  

2005 Cry1Ac Results

μg/ml Cry1Ac (100% corrected mortality).  Only one Arizona population
was tested at the 1.0 µg Cry1Ac/ml dose and had a corrected mortality
of 66.9%.  The susceptible laboratory colony used as a control group
showed 100% mortality to the test concentration of 10 µg Cry1Ac/ml. 
Overall, the authors concluded that PBW remains susceptible to Cry1Ac
and that are no indications of resistance in the field.  The 2005
results from Arizona are summarized and compared with historical data in
Table 6 below.

2005 Cry2Ab2 Results

	

 1 μg/ml and 10 μg/ml Cry2Ab2.  The susceptible laboratory colony used
as control had 100% mortality at the 10.0 µg Cry2Ab2/ml concentration. 
As with Cry1Ac, the authors concluded that the sampled PBW populations
remained highly susceptible to Cry2Ab2.  The 2005 Arizona data are
summarized in Table 7 below.

2005 Field Efficacy Studies

In addition to the susceptibility bioassays, the Arizona monitoring
group sampled large numbers of Bt and non-Bt cotton bolls throughout the
state (obtained from 40 pairs of Bt and non-Bt fields).  The procedures
were similar to the boll sampling that was also conducted during 2004. 
Infestation rates in the 2005 field efficacy study were 0.37% average
infested Bt bolls (range not reported in Dennehy et. al, 2006 as this
was a preliminary report).  This rate is slightly higher than that
reports for the 2004 growing season.  The average infestation Bt cotton
rate for 2004 was 0.34% bolls and for 2003 was 0.21% (range 0 to 1.40%)
bolls.   For non-Bt cotton, 8,100 non-Bt bolls were examined with an
average infestation rate of 24.0% (range not reported) compared with
21.7% (range from 0 to 100%) in 2004 and 29.0% (range from 0 to 100%) in
2003).  In 2004, subsequent analysis of the Bt bolls determined that
many were non-expressing off-types (Dennehy et al., 2005).  No analysis
was provided for the 2005 data (Dennehy et al., 2006 is a preliminary
report).  It is likely that many of the collected Bt bolls with PBW
larvae will be non-Bt expressing off-types rather than the result of
adaptation to Bt toxins (see Dennehy et al., 2005 for discussion of 2004
results).  Attention should still be given to any increase in boll
infestation rates in Bt fields. Results of the field efficacy studies
conducted in Arizona from 1995 to 2004 are shown in Figure 7 (at the end
of this review).

Table 6. Field-Collected PBW Mortality to Discriminating Concentrations
of Cry1Ac from 1997 to 2005 (Reynolds, 2006; updated with Dennehy et al.
2006) 

Year	Average Mortality of Field Collected PBW (%)1

	1.0 µg Cry1Ac/ml concentration 	10 µg Cry1Ac/ml concentration

1997	57.4	94.1

1998	90.6	99.9

1999	97.9	100

2000	97.4	100

2001	94.8	99.4

2002	85.7	99.8

2003	68.3	99.8

2004	95.4	99.9

2005	66.92	100.0

1 Mortality values are corrected for mortality observed in control
groups.

2  Only one location was tested at this test concentration.

Table 7. Field-Collected PBW Mortality to Discriminating Concentrations
of Cry2Ab2 from 2003 to 2005 (Reynolds, 2006; updated with Dennehy et
al., 2006) 

	Year		Average Mortality of Field Collected PBW (%)1

		1.0 µg Cry2Ab2/ml dose 		10 µg Cry2Ab2/ml dose

2003		97.3		99.9

2004	99.1	100

2005	100	100

1 Mortality values are corrected for mortality observed in control
groups.

Discussion

Since the PBW monitoring methodology has remained consistent throughout
the resistance monitoring program, the data can be placed in a
historical context to evaluate long-term shifts in susceptibility.  The
10.0 µg/ml concentration for both Cry1Ac and Cry2Ab2 is essentially a
true discriminating dose, i.e. a PBW LC99 that can be used to
distinguish potentially resistant insects from susceptible ones.  

Through 2005, ten years of monitoring data have now been tabulated for
Cry1Ac and three years for Cry2Ab2.  Based on BPPD’s analysis of the
larval susceptibility data and boll infestation data, PBW susceptibility
to both toxins remains high (see Tables 6 and 7 above) and there are low
boll infestation rates in Bt fields as observed in the field efficacy
trials.  Monitoring data and field efficacy data from 1997 to 2005
indicate there hasn’t been an increase in resistance to Cry1Ac and to
Bt cotton (see Dennehy et al., 2002, 2003, 2004, 2005, 2006; Unnithan et
al., 2004).  Tabashnik et al. (2005a) confirm that the resistance allele
frequency did not increase over the period from 1997 to 2004 based on
bioassays.   

A resistance allele frequency of 0.16 was estimated for recessive
resistance to Cry1Ac based on bioassay results from collections of pink
bollworm at 10 sites in Arizona in 1997 (Tabashnik et al., 2000).  
Subsequent work at the University of Arizona showed that resistant
larvae from at least four different laboratory-selected strains survived
on Bt cotton, resistance was recessive and tightly linked with mutations
in a cadherin gene that encodes a Cry1Ac-binding protein.   Three
cadherin resistance alleles were identified (r1, r2, and, r3). 
University of Arizona researchers do not know why the frequency of
resistance was so high in 1997.  Setting aside this unusually high
frequency of pink bollworm resistance to Cry1Ac detected in 1997, the
bioassay results since 1998 provide evidence that pink bollworm
resistance to Bt  cotton did not increase substantially from 1998 to
2005.  Indeed, no resistant individuals were detected in bioassays of
5,358 individuals sampled from Arizona and California in 2005. 
Further, no alleles for resistance were detected in 5,571 individuals
sampled from Arizona, California, and Texas from 2001 to 2005 (Tabashnik
et al., 2006).  Therefore, the resistance allele frequency of 0.01 used
as one of the parameter values in the simulation model is quite
conservative and is much higher than the gene frequency that was
detected in 2005 (see modeling section below). 

Field efficacy data indicates that there has been a numerical, but not a
statistically significant, increase in percent infested bolls in Bt
cotton from 2003 to 2005 (data from 1995 to 2004 shown in Figure 7;
Dennehy et al., 2006 for 2005 data).  Tests of infested bolls (n=35)
collected from Bt fields in 2004 revealed that a large portion of
infested bolls (over 90%) did not have Cry1Ac protein in the seeds. 
Similar analyses have net yet been performed for Bt bolls collected in
2005 (Dennehy et al., 2006).   Infestation levels in Bt fields have
averaged ≤0.370% over the past ten years; an amount of less than four
pink bollworms per 1000 bolls (Figure 7).   

The ACRPC has indicated that follow-up susceptibility testing will be
conducted with PBW larvae recovered from Bt bolls (verified expressing
the Bt toxin).  This is important because Bollgard and Bollgard II are
considered high dose for the control of PBW; therefore, larvae recovered
from Bt bolls may be heterozygous or homozygous for Bt toxin
resistance.  The determination that these larvae are carrying heritable
resistance traits could provide an early indication of a resistance
problem.  Under the PBW eradication program, in the unlikely event that
widespread field resistance be a concern, additional actions as
prescribed by the Remedial Action Plan for PBW Resistance to Bt Cotton
(see Appendix 1) would be implemented.  Follow-up testing will also be
conducted on survivors of the 10 µg/ml Cry1Ac and Cry2Ab2
discriminating concentrations.  These larvae may be homozygous for
Cry1Ac or Cry2Ab2 resistance alleles and warrant additional scrutiny.  

There have been pink bollworm survivors at the 10 µg Cry1Ac/ml
discriminating concentration in 2001, 2002, 2003, and 2004 (Figure 8).  
Given that some PBW have survived the 10 µg Cry1Ac/ml discriminating
concentration in previous years, it is possible that resistance alleles
are relatively common in PBW populations in western cotton growing
regions. However, selection data and PCR data don’t support this
interpretation (see Tabashnik et al., 2006).   Survivors may not be
genetically-resistant. For example, in 2001 there were 31 survivors from
Arizona at the diagnostic concentration in bioassays (Tabashnik et al.,
2005a), yet selection with Cry1Ac in diet did not yield a resistant
strain.  This lack of response to selection suggests that the survival
was not heritable and thus the 2001 resistance allele frequency was
overestimated from bioassays.  In contrast, with strains derived from
Arizona cotton fields in 1997, as few as three generations of selection
with Cry1Ac produced strains capable of surviving on Bt cotton
(Tabashnik et al. 2000, 2005b).  Contamination of field-derived strains
by resistant laboratory strains may have been possible for bioassays
conducted during 2001 to 2004, although unlikely, according to the
explanation provided in Tabashnik et al. (2006).  There were no
resistant strains available to contaminate the bioassays in 1997.  Also,
no such contamination affected bioassay results in 1999, 2000 and 2005
when no resistant individuals were detected in field-derived strains
(total n = 11,400 larvae tested at the diagnostic concentration).  One
final piece of information that should be considered is that researchers
in Arizona have not been able to select for a resistant strain from
field collections from Arizona despite trying every year.  Only in 1997,
were resistance colonies established from field collections (see
Tabashnik et al., 2005a).

Considering the high use of Bt cotton (93% in the eradication zone in
2006; 100% was allowable for 2006) under the four-year PBW eradication
program, it will be important to closely monitor PBW and Bt cotton for
resistance to Cry1Ac and Cry2Ab2 and possible unexpected field damage.

Simulation Modeling Comparing Population Suppression Vs. Resistance Risk
for the Duration of Eradication Program  

The Agency required that simulation modeling be used to compare the
impact of population suppression vs. resistance risk over the period of
the eradication program.   The simulation model used was a revised
version of the spatially explicit, stochastic model discussed in
Sisterson et al. (2004).  The simulations examined population
suppression (number of PBW per ha) and risk of resistance to Bt cotton
(rate of increase of resistance allele frequency).  This model considers
resistance controlled by a single, recessive gene.  This model is based
on PBW resistance to Cry1Ac.  Resistance to Cry2Ab2 is not considered in
the model.  Table 8 below provides the parameter values for the
simulation model.  Default values are based on best estimates available
in the public literature and recent field data.  Recent PBW data have
been collected in Arizona regarding the genetic basis of resistance to
Bt cotton that produces Cry1Ac, the frequency of resistance alleles in
field populations, population sizes, population dynamics, and
movement.  Modifications to the model include the release of sterile
moths.   Default values are based on best estimates available in the
public literature.  Alternatives to the default values were
systematically evaluated for parameters such as resistance allele
frequency and movement.   A variety of scenarios were simulated using
the best estimates of the parameter values as well as more optimistic
and more pessimistic scenarios.   Relative to simulations using best
estimates for parameters, more optimistic scenarios might underestimate
the risk of resistance, while more pessimistic scenarios might
overestimate the risk of resistance.   Modeling will also explore the
impact on population suppression and resistance risk of alternative
management options (e.g., variable refuge size and release rates of
sterile moths).  A region with 4096 cotton fields was modeled for four
years (the extent of the intended PBW eradication program in Arizona). 
 Overwintering larvae from the fourth year are checked to assess the
following criteria.

Resistance will occur if the resistance allele frequency exceeds 0.50.

Population recovery will occur if the final population is equal to or
greater than the initial population size (29,000 overwintering larvae
per field is the default value).

Population suppression will occur if the mean PBW density in the region
is equal to or less than 0.1 overwintering larvae per 15 ha (=0.0067
larvae per ha).

Regional loss will occur if all fields in the region modeled have 0 PBW.

Results

In the analyses to date, 12 sets of assumptions, each of them simulated
5-16 times, for a total of 128 simulation runs were examined.  In all
cases, except for the default case, the initial Cry1Ac resistance allele
frequency (r) was 0.01, which is ten times higher than the default value
of 0.001.  This is a realistic estimate based on the bioassay data
discussed in Tabashnik et al., 2005a (and above in this review) as well
as DNA screening (Tabashnik et al., 2006).  In all cases, 500 sterile
moths per ha were released in non-Bt cotton fields, with one release per
three days.  Results of the simulation outcomes are summarized in Table
9.  In 11 of 12 sets of assumptions examined, the simulated eradication
program eliminated the PBW from the 4096 fields modeled in two years or
less without the development of resistance.   These included
“worst-case or pessimistic” simulations in which resistance was
inherited as a dominant trait, 90% Bt cotton and 10% non-Bt cotton
refuges (rater than 100% Bt cotton), no fitness cost of resistance
(rather than a 10% fitness cost in homozygous resistant insects, and
release rates of 1, 2, 3, 4, 5, 10, or 15 sterile moths per ha in Bt
cotton fields (rather than 75 sterile moths per ha).   Tabashnik
(personal communication, B. Tabashnik, U. of Arizona to S. Matten,
USEPA/OPP/BPPD, dated September 14, 2006) noted that more recent
simulations show regional loss (i.e., 0 PBW) with 2 per ha per release
in Bt cotton and 10 per ha per release in non-Bt cotton. 

In the one exception, PBW was not removed from the region when there was
no release of sterile moths in Bt fields, 90% Bt cotton, and r = 0.01 in
all five replications.  In this case, the population density declined by
98% (460 final overwintering larvae per field/29,000 starting
overwintering larvae per field) and the resistance allele frequency
increased from 0.01 to 0.02 after four years.  While the resistance
allele frequency doubled after four years, it is still far below the
0.50 value typically used as a criterion for a resistance problem.



Table 8. Parameter values for eradication model (revised from Sisterson
et al., 2004).

Default values, which are used unless noted otherwise for parameters
with more than one value, are indicated by an asterisk. [Taken from
Volume 5, MRID# 469048-05]

Parameters							Values

Adults							

Mean % of adults that leave their natal field 			10, 55*, 75 

Number of eggs per female per day in Bt cotton fields 	10 

Number of eggs per female per day in non-Bt cotton fields 	10 

Mean % of adults that die each day 				10 

Egg-pupae 

Mutation rate (from S to R per allele) 			5 x 10-5

Mean % of SS and RS killed in non-Bt cotton fields 	79.2

Mean % of RR killed in non-Bt cotton fields 		79.2, 81.3*(10% fitness
cost)

Mean % of SS and RS killed in Bt cotton fields 		99.8a, 100*

Mean % of RR killed in Bt cotton fields 			79.2, 83.2*(incomplete R=0.9)

Development time (degree days) 				433

Mean % of larvae that die during overwintering 		95

Region

Initial R allele frequency 					0.0001, 0.001*, 0.01

Number of fields 						4096 (64 X 64 square)

Size of fields 							15 hectares

Percentage of Bt fields 					80, 85, 90, 95, 100*

Percentage of Bt plants in Bt fields 				99a, 100*

Distribution of fields 						Random

Carrying capacity per field 					4,200,000

Initial overwintering larvae per field 				2900, 29,000*, 290,000

a 99.8% mortality of RS and SS simulates 100% Bt fields that have 99% Bt
cotton plants and 1% non-Bt cotton plants (contaminants); 100% die on
the Bt plants, 79.2% die on the non-Bt plants (0.99 X 100% + 0.01 X
79.2% = 99.8%) 



Steriles 

Release period 

Frequency of releases in each field 

Sex ratio of steriles 

Steriles per ha per release in Bt cotton fields 

Steriles per ha per release in non-Bt cotton fields 

May 1-Oct 15 (1st bloom to defoliation) 

1 per 3 days per field 

1 female: 1 male 

0, 1, 2, 3, 4, 5, 10, 15, 75* 

0, 100, 500*, 1000 

Pheromone ropes only in non-Bt cotton fields

All non-Bt fields treated once early in season 

May 17-June 20 (6-leaf stage) 

Daily % reduction in fecundity caused by 

pheromone ropes 					  20, 40*, 60 for 30 days 

Insecticide & pheromone sprays only in non-Bt cotton fields 

Spray threshold 					>60 no spray

(check sterile male:native male ratio weekly)	30-59 spray pheromone

							0-29 spray pheromone + insecticide

Daily reduction in fecundity caused by pheromone sprays 	20, 40*, 60 for
14 days

Mean % of adults killed daily by insecticide			37 per day for 5 days

Larvae are not killed by sprays				95 per day for 5 days



Table 9. Simulation outcomes: effects of pessimistic assumptions on
simulated outcomes of the pink bollworm eradication program in Arizona.
In all cases except the default case, the initial resistance allele
frequency (r) was 0.01, which is ten times higher than the default value
of 0.001.  In all cases, 500 sterile moths per ha were released in
non-Bt cotton fields, with one release per three days. [Taken from
Volume 5, MRID# 469048-05] 

Parameter values different from default values 			Outcome* 

None (all parameters at default values) 				Loss in 1 year 

Initial resistance allele frequency (r) = 0.01 				Loss in 2 years 

No fitness cost, r = 0.01 						Loss in 2 years 

Dominant resistance to Bt cotton, r = 0.01 				Loss in 2 years 

90% Bt cotton, r = 0.01 						Loss in 2 years 

Steriles released in Bt cotton fields 

at 1, 2, 3, 4, 5, 10, or 15 steriles per ha 

90% Bt cotton, r = 0.01 						averages after 4 years 

No steriles released in Bt cotton fields 				r = 0.02 

                                                                        
                                   460 larvae per field

                                                                        
                                                          (overwintering
survivors)

*loss means that no pink bollworm were present in any of the 4096 cotton
fields modeled 

 

Discussion

Even with pessimistic parameter assumptions, simulations of the
four-year eradication program in Arizona yielded suppression of pink
bollworm without creating a severe problem with resistance developing to
Bt cotton.  The current simulations suggest that the release of sterile
moths in Bt cotton fields is important for driving PBW population
densities to extremely low levels.  However, these are preliminary
simulations and certain parameter values are best estimates rather than
actual field measurements.  Additional modeling simulations should be
done using field values.  This will allow partial field-validation of
the model.  Dr. Bruce Tabashnik (University of Arizona) has indicated
that the data from the field will be used to test the model’s
predictions about the ratios of sterile:native moths and resistance
allele frequency (e-mail from B. Tabashnik, University of Arizona to S.
Matten, USEPA/OPP/BPPD, dated September 14, 2006).  If the model’s
predictions are wrong, Tabashnik notes that any incorrect assumptions
will be changed to improve the accuracy of the model.   The results of
the additional modeling simulations should be submitted to the Agency
for review as a supplemental submission.

Tabashnik (personal communication, B. Tabashnik, U. of Arizona to S.
Matten, USEPA/OPP/BPPD, dated September 14, 2006) writes that additional
pessimistic assumptions will be investigated to determine which
conditions might cause severe resistance problems.  Some of these
simulations are currently being performed. Some of assumptions were
modified and another set of scenarios using sensitivity analysis varying
emigration (10, 55, or 75%), dominance (recessive or dominant), number
of fields in the region (400 or 4096), percentage of Bt fields (80, 95,
100), carrying capacity per field (2,885,  28,885, 288,850), and numbers
of steriles per ha per release in Bt and non-Bt cotton fields (0,0;
2,10; 100, 500; and 200, 1000) were run.  As in the first 12 scenarios
run (i.e., the results in Vol. 5, MRID# 469048-05), all of the
additional scenarios yielded regional loss (i.e., no PBW) in 2 years
except for one case.  The only case in which loss did not occur was with
no steriles released in Bt or non-Bt cotton (0, 0).  The lowest non-zero
release rate tested was 2 per ha per release in Bt cotton and 10 per ha
per release in non-Bt cotton.  This low release rate scenario yielded
regional loss in 2 years.    Based on the actual sterile release rates
(thru August 25, 2006; see Table 4 above), and the actual percentage of
Bt and non-Bt cotton fields see Table 1 above), the range of input
parameters used in the modeling simulations was reasonable (see Table
8).  New simulations using the field data collected in 2006 should be
used to confirm the outcomes of the preliminary modeling.

Pink bollworm resistance to the Cry2Ab2 toxin (present in Bollgard II)
is not considered in the modeling.  The use of 100% Bollgard and
Bollgard II both have been authorized for use under the two 24(c)
registrations.  Based on sales information through the 2005 growing
season, the ratio of Bollgard to Bollgard II cotton sales in Arizona is
approximately 95:5.  This means that there has been very little
selection pressure to the Cry2Ab2 toxin, to date, in Arizona.  The
greatest selection pressure for resistance is to the Cry1Ac toxin
present in Bollgard and Bollgard II cotton.  Sales information for the
2006 growing season will be provided to the Agency by Monsanto in
January 2006 (requirement of the terms and conditions of the Bollgard
and Bollgard II registrations).  No information was provided by the
state of Arizona as to the breakdown of Bollgard and Bollgard II cotton
use in the eradication zone (4626 fields mapped) during the 2006 growing
season.   What is known is that there were approximately 93% Bt cotton
fields and 7% non-Bt cotton fields in the eradication zone for 2006.  

If some or all of Arizona’s Bt cotton had two toxins, Cry1Ac + Cry2Ab,
evolution of resistance would be much less likely than it is with only
Cry1Ac.  Modeling resistance to cotton that produces only Cry1Ac is the
most pessimistic.  The modeling predictions (using only Cry1Ac
resistance), therefore, are conservative, i.e., they tend to
overestimate resistance risk.  Several researchers have modeled the
benefit of managing resistance evolution to two toxins with dissimilar
modes of action using a pyramided approach (e.g., Zhao et al. 2005;
Roush 1998).

  



References

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Carrière, and B. Tabashnik, 2002.  Monitoring for Cry1Ac protein
susceptibility among field populations of pink bollworm during the 2001
cotton growing season.   Cooperative Extension. The University of
Arizona.  Extension Arthropod Resistance Management Laboratory.
Unpublished study submitted to EPA.  MRID #  457065-01.

Dennehy, T., S. Brink, B. Wood, D. Holley, G. Unnithan, Y. Carrière,
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Dennehy, T., G. Unnithan, S. Brink, B. Wood, Y. Carrière, B. Tabashnik,
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Dennehy, T., G. Unnithan, S. Brink, B. Wood, Y. Carrière, and B.
Tabashnik, 2005.  Susceptibility of Bt toxins Cry1Ac and Cry2Ab2 to
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Dennehy, T. J., G. C. Unnithan, S. Brink, and B. Wood.  2006. 
Preliminary Report of susceptibility to Bt toxins Cry1Ac and Cry2Ab2 of
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Unnithan, G.C., S. Brink, B. Wood, Y. Carrière, B. Tabashnik, L.
Antilla, and M. Whitlow.  2004.  Susceptibility of Southwestern pink
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EPA Reviews of the Resistance Monitoring Data

U.S.  Environmental Protection Agency (U.S. EPA).  2001.  Bt
plant-incorporated protectants October 15, 2001 biopesticides
registration action document.  Available at
http://www.epa.gov/pesticides/biopesticides/pips/bt_htm.

Matten, S. 2003.  Technical review of Monsanto’s submissions:
resistance monitoring protocols for monitoring of Cry1Ac susceptibility
among field populations of pink bollworm, tobacco budworm, and cotton
bollworm.  S. Matten memorandum to L. Cole, September, 2, 2003.

Reynolds, A.  2004b.  Review of 2001-2 monitoring data submitted by
Monsanto for Bt cotton (Bollgard).  A. Reynolds memorandum to L. Cole,
September 8, 2004.

Reynolds, A.  2004c.  Review of preliminary monitoring data submitted by
Monsanto for Bollgard II Bt cotton.  A. Reynolds memorandum to L. Cole,
May 26, 2004.

Reynolds, A.  2005.  Review of 2003 monitoring data submitted by
Monsanto for Bt cotton (Bollgard and Bollgard II).  A. Reynolds
memorandum to L. Cole, June 8, 2005.

Reynolds, A. 2006.  Review of 2004 pink bollworm monitoring data and
revised monitoring protocol submitted by Monsanto for Bt cotton
(Bollgard and Bollgard II).  A. Reynolds memorandum to L. Cole, July 20,
2006.

Other Citations

Bartels, D., M. Walters.  2006.  Spatial analysis of native and sterile
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22, 2006.  

Carrière, Y., C. Ellers-Kirk, M. Sisterson, L. Antilla, M. Whitlow,
T.J. Dennehy and B.E. Tabashnik.  2003.  Long-term regional suppression
of pink bollworm by Bacillus thuringiensis  cotton.  Proc. of the
National Academy of Sciences.  USA 100:  1519-1523.

Carrière, Y., M. E. Nyboer, C. Ellers-Kirk, J. Sollome, N. Colletto, L.
Antilla, T. J. 

Dennehy, R. T. Staten and B. E. Tabashnik. 2006. Effect of resistance to
Bt cotton on pink bollworm (Lepidoptera: Gelechiidae) response to sex
pheromone. J. Econ. Enomol. 99:  946-952.

Gahan, L. J., F. Gould, and D. G. Heckel. 2001. Identification of a gene
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resistance in Heliothis virescens. Science 293: 857-860. 

Henneberry, T.J. and S.E. Naranjo.  1998.  Integrated management
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Keaveny, D., J. Hessler, J. Rudig.  2006.  Local movement of sterile
pink bollworm moths in the San Joaquin Valley of California.  Special
Report 2006.  

Matten, S.R.  2005.  Technical review of Monsanto’s sales and acreage
data for Bollgard® and Bollgard II® Cotton as well as reports of
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November 9, 2005.

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compared with natives.  J. of Econ. Entomol. 87:  680-686.

Morin, S.R., W. Biggs, M.S. Sisterson, L. Shriver, C. Ellers-Kirk, D.
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Dennehy, J.K. Brown, and B.E. Tabashnik.  2003.  Three cadherin alleles
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Morin, S., S. Henderson, J. A. Fabrick, Y. Carrière, T. J. Dennehy, J.
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1225-1233. 

Reynolds, A.H. 2004a.  Review of Monsanto’s proposed amendment to
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Gelechiidae) males in transgenic cotton that produces a Bacillus
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Tabashnik, B. E., Y.-B. Liu, S. Morin, D. Unnithan, Y. Carrière, and T.
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Tabashnik, B. E., R. W. Biggs, D. M. Higginson, S. Henderson, D. 
Unnithan,  C. Ellers-Kirk, M. S. Sisterson, T. J. Dennehy, Y. Carrière,
et al. 2005a. Association between resistance to Bt cotton and cadherin
genotype in pink bollworm. J. Econ. Entomol. 98: 635-644. 

Tabashnik, B. E., T. J. Dennehy, and Y. Carrière. 2005b. Delayed
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U.S.A. 43: 15389-15393. 

Tabashnik, B.E., J.A. Fabrick, S. Henderson, R.W. Biggs, C.M. Yafuso,
M.E. Nyboer, N.M. Manhardt, L.A. Coughlin, J. Sollome, Y. Carrière,
T.J. Dennehy, and S. Morin.  2006. DNA screening reveals pink bollworm
resistance to Bt cotton remains rare after a decade of exposure. J.
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Xu, X., L. Yu and Y. Wu. 2005. Disruption of a cadherin gene associated
with re楳瑳湡散琠⁯

Cry1Ac δ-endotoxin of Bacillus thuringiensis in Helicoverpa armigera.
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Microbiol. 71: 948-954. 

Zhao, J.-Z., J. Cao, H.L. Collins, S.L. Bates, R.T. Roush, E.D. Earle
and A.M. Shelton,  2005.  Concurrent use of transgenic plants expressing
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FIGURES

Figures 1-4.  Arizona Cotton Research Protection Council
copywrite-protected Bt/non-Bt field maps for Central and Eastern Arizona
and the sterile moth release zone are in a separate attachment (Appendix
4).

Figure 5.  Scheme:  Suggested Use of Trapping and Map Data

Figures 6A-H.  Kriging Maps of the Sterile and Native Pink Bollworm
Trapping Data

Figure 7.  Field efficacy to Bt cotton in Arizona: 1995 to 2004

Figure 8.  Changes in pink bollworm susceptibility to Cry1Ac in Arizona
from 1997 to 2004

Figure 5.  Scheme:  sUGESSTED USE OF trapping and map data submitted as
supporting documentation FOR

SPECIAL LOCAL NEED REGISTRATION 24c, FOR bt COTTON

 FOR THE PURPOSE OF PINK BOLLWORM ERADICATION

 

 

 

                   

 

Figure 6A.  Kriging map of Arizona Sterile Pink Bollworm Trapping Data 
- Week of June 25, 2006.

Figure 6B.  Kriging map of Arizona Sterile Pink Bollworm Trapping Data 
- Week of July 2, 2006.

Figure 6C.  Kriging map of Arizona Sterile Pink Bollworm Trapping Data 
- Week of July 9, 2006.

Figure 6D.  Kriging map of Arizona Sterile Pink Bollworm Trapping Data -
Week of July 16, 2006.

Figure 6E.  Kriging map of Arizona Native Pink Bollworm Trapping Data  -
Week of June 25, 2006.

Figure 6F.  Kriging map of Arizona Native Pink Bollworm Trapping Data  -
Week of July 2, 2006.

Figure 6G.  Kriging map of Arizona Native Pink Bollworm Trapping Data  -
Week of July 9, 2006.

Figure 6H.  Kriging map of Arizona Native Pink Bollworm Trapping Data  -
Week of July 16, 2006.

Figure 7.  Field efficacy of Bt cotton in Arizona:  1995 to 2004.  Data
from 1995 to 1997 were reported by Flint et al.(1995) and Flint and Park
(1996).  All other data were collected by the Arizona Cotton Research
and Protection Council.  Shown are means of the percent boll infestation
(bolls with ≥ 3rd instar PBW) for pairs of Bt  cotton (left axis) and
non-Bt cotton fields (right axis) sampled each year from 1995 to 2004. 
The number of pairs of Bt  and non-Bt fields (N) is indicated for each
year.  [Figure from Dennehy et al., 2005]

bioassays of 1.0 and 10 μg Cry1Ac/ml diet of field collections made
throughout Arizona in 1997 (n=9), 1998 (n=12), 1999 (n=14), 2000 (n=17),
2002 (n=13), 2003 (n=16), and 2004 (n=13).   No larvae from any tests of
2005 strains survived treatments of 10 μg Cry1Ac/ml diet. [Taken from
Dennehy et al., 2006]



APPENDICES

Appendix 1:  A Remedial Action Plan to Respond to Pink Bollworm
Resistance to Bt Cotton in Arizona

Appendix 2:  Pink Bollworm Trapping and Sampling Methodology

Appendix 3:  Tabashnik’s Answers, dated 9/14/06, to Matten’s
Questions, dated 9/8/06, Re: PCR 

Appendix 4:  Arizona Cotton Research Protection Council copy
write-protected field maps (separate document)



APPENDIX 2  Pink Bollworm Trapping and Sampling Methodology [Taken from
letter dated September 14, 2006 from L. Antilla, Arizona Cotton Research
and Protection Council to S. Matten, USEPA/OPP/BPPD]

TRAP DENSITY

NON-BT COTTON

1 trap per ten acres or 1 trap per field on fields less than ten acres.

BT

1 trap per forty acres on small contiguous blocks of fields with out
biological separation.

1 trap per eighty acres on large contiguous blocks of fields with out
biological separation.

TRAP PLACEMENT

Protocol calls for traps to be placed at or near the northeast corner of
the field in a protected location (near a permanent structure such as a
telephone pole). If a field has more than one trap, the traps are evenly
spaced and numbered in a counter clockwise manner. Traps are placed as
near to the field edge as possible while not obstructing the movement of
equipment in and out of the field. Traps are attached to a wooden survey
stake in order to maintain traps at canopy height.

TRAP SERVICE

Traps are regularly serviced once each week unless environmental
conditions are prohibitive i.e. moisture soaked terrain inaccessible by
4 wheel drive. When a trap is serviced the entire trap is removed and a
new trap with a new Pink Bollworm pheromone lure is placed in the trap.
The “old” trap is labeled with the service date and the crop stage.
Trap locations are bar-coded and each “new” trap is labeled with the
field number and trap number. Traps are baited a with rubber dispenser
impregnated with a 4 mg dose of Shinitsu Corporation Hexadecadienyl
acetate (Gossyplure, Pheromone) that has been subjected to field
bioassay for field activity by USDA personnel.

The traps removed from the field are transported to the field offices in
a protected manor for identification.

MOTH IDENTIFICATION

Traps are brought in from the field each week; identifiers stationed at
each field office inspect each trap. All Pink Bollworm moths are counted
and recorded as either “native” or “sterile”. The sterile pink
bollworms are dyed red through the media they are fed in the rearing
facility. The trapping date (date the trap was removed from the field),
field number, trap number and crop stage are all recorded along the
“native” and “sterile” counts for entry into the database. Any
questionable determinations regarding the identity of moths are
forwarded to the principle identifier. 

At the discretion of program management and as readily available samples
for Dr. Tim Dennehy’s Extension Arthropod Resistance Management
Laboratory (EARML), moths are saved for testing related to BT resistance
or genetic identity. The moths collected for these functions are placed
in alcohol and kept in a freezer to protect from deterioration until
testing can be completed.

BOLL SAMPLING

Boll sampling is conducted on randomly selected fields or pairs of
fields throughout the program area to determine representative
infestation levels. When possible, a BT and a NON-BT field in close
proximity at the randomly selected location are included. The purpose of
this exercise is to approximate the approach taken in Arizona since 1998
wherein, randomly selected paired BT and NON-BT fields were intensively
sampled late season for comparative infestation levels and resistance
monitoring. 

Based on trapping information, history and targeted field inspection,
other fields are checked for boll infestation as needed. Program
personnel make these decisions by dedicating the majority of our
resources on field based activities whereby anomalies are isolated and
investigated to the benefit of the producers in the program. Due to the
targeted approach in this instance any findings are neither
representative nor random and therefore statistically not indicative of
program wide infestation levels.

Sampling each of the four thousand six hundred and twenty six fields
each week is not logistically or economically feasible and would
certainly be undesirable to producers in the program.

 

The initial boll survey sampling was performed for 4 sampling cycles (1
sample every other week). Additional sampling will be conducted using
boll boxes. Boll boxes data is much more reliable as bolls are picked
and then placed in boxes (cages) and stored at controlled temperatures
until any organisms inside the bolls have emerged and can be counted.

STERILE/NATIVE TRAPPING DATA

By statute, all growers within the program area must report NON-BT
cotton in a timely manner. BT cotton reporting is not legally required
and therefore must be ground proofed by program personnel. Circumstances
do exist wherein late planted or unreported cotton results in data
beginning the week after traps are deployed.

Trapping data is not a self contained gauge of populations. Pheromone
traps are subject to hindrance from many biological, environmental and
seasonal influences. Inherent variability in trapping information from
specific trap location data must be evaluated as an aggregate from
multiple data points to be meaningful per the maps provided by Dave
Bartel and the statistical data model. Moth numbers found in traps are
not directly proportionate to release levels on that field or the
surrounding area. Native populations in the program area are still very
high. Native and Sterile females produce pheromone which competes with
the traps. As the native populations become less significant the
pheromone traps will become more consistent population indicators as
individual data points. As discussed in the trapping section above,
there are periodic instance where traps cannot be accessed, in these
instances no data is available. 

As indicated in the trapping section of this document, BT fields are
grouped together in regards to trap allocation. Not every BT field has a
trap directly assigned to it however; traps are distributed through the
region to produce a representative sample of the region and all of the
fields therein. Trapping each BT cotton field individually is cost
prohibitive and logistically impossible within the constraints of the
Pink Bollworm Eradication Program. This approach is in strict contrast
to non eradication program grower practices where BT cotton is not
monitored with pheromone traps. 

All fields have received sterile release every week on timetable with
minor variations due to weather and or chemical treatment. Chemical
treatment only affects sterile release on NON-BT cotton as BT cotton is
not treated with chemical or pheromone treatments. Sterile release has
been unremitting once sterile release began. No release days have been
compromised due to mechanical failure, moth supply or due diligence.
Sterile moths in excess of 1,137,012,553 have been released over Arizona
BT and NON-BT cotton as of September 5, 2006 within the Pink Bollworm
Eradication Program.”



APPENDIX 3: Tabashnik’s Answers, dated 9/14/06, to Matten’s
Questions, dated 9/8/06, Re: PCR  [Taken from e-mail, B. Tabashnik, U.
of Arizona to S. Matten, USEPA/OPP/BPPD, dated September 14, 2006]

1.  Is the Molecular analysis method in the submission to EPA, the one
published in Tabashnik et al. 2006?  If so, I need an e-mail back to me
identifying that this is indeed the method used.

Yes.  The molecular analysis method in the submission to EPA is the one
published in Tabashnik et al. 2006.  Key portions of the paper
describing the method are provided below.  Please note that initial
tests use only a small portion of each field-sampled individual
(Tabashnik et al. 2005b), so that re-testing of individuals is possible
if desired.

INTRODUCTION

“In laboratory-selected strains of pink bollworm and at least two
other major lepidopteran pests of cotton, mutations in a cadherin gene
are tightly linked with recessive resistance to Cry1Ac (Gahan et al.
2001, Morin et al. 2003, Xu et al. 2005).  In several
laboratory-selected strains of pink bollworm, three mutant alleles (r1,
r2, and r3) of a cadherin gene (BtR) are associated with resistance to
Cry1Ac and survival on Bt cotton (Morin et al. 2003, 2004; Tabashnik et
al. 2004, 2005b).  Each r allele has a deletion predicted to eliminate
at least eight amino acids upstream of the putative Cry1Ac-binding
region of cadherin protein (Morin et al. 2003).  We previously developed
a PCR-based method for detecting the r1, r2 and r3 alleles in pink
bollworm (Morin et al. 2004).  We isolated, cloned and sequenced the
genomic region spanning the mutation in each r allele and designed
allele-specific PCR primers for each region.  The method can detect any
of the three r alleles in a single heterozygote (r1s, r2s, or r3s)
pooled with DNA from the equivalent of 19 susceptible (ss) individuals
(Morin et al. 2004).”

METHODS:  “DNA Preparation and PCR.  Insects collected from bolls and
traps were stored in ethanol at –20oC.  DNA was extracted using DNAzol
(Tabashnik et al. 2005b) and PCR was done as described by Morin et al.
(2004).  The maximum number of individuals tested per pool was 5 for
samples from 2001-2003 and 11 for samples from 2004-2005.”

2.  The late season sampling method developed by Dennehy and Tabashnik--
I don't have the specific protocol.  Please send it or perhaps it is in
the Tabashnik et al. 2006 manuscript. I see a couple of sentences
describing the plan, is this it?  Similar to Tabashnik et al. (2006),
details coming soon.

3.  What is the method for estimating false negatives? false positives?
What is the likelihood of non-detection?  Please clarify.

The methods for estimating false negatives, false positives, and the
likelihood of non-detection are detailed in Morin et al. (2004) and
Tabashnik et al. (2006), as well as below:

A. False negatives.  

False negatives are possible from three causes: i) The PCR reaction is
not working properly, ii) The cadherin DNA of field-sampled insects is
not amplified, iii) The PCR is working and cadherin DNA is amplified
from field samples, yet r alleles are present and are not detected.

i) To determine if the PCR reaction is working properly, we use known
positive controls in every set of samples tested.  This is a standard
method.  Known positive controls are samples of DNA from our
laboratory-reared strains that contain r alleles, which are known to the
person running the PCR reaction.  For example, every test of
field-sampled insects for the presence of an r1 allele includes a gel
lane in which DNA from one or more laboratory-reared individuals with
the r1 allele is run simultaneously with the field samples.  

If the known sample of r1 DNA does not yield a positive result for r1,
the test of the field sample is not valid and must be repeated.  In this
case, PCR reaction conditions are corrected until the known controls
yield positive results with the simultaneously tested field samples. 
Such corrections usually involve systematic replacement of reagents
(primers, Taq, etc.) to ensure all are working properly.  Because only
tests yielding positive results for known positive controls are included
in our analysis of the data, this source of false positives has an
effective rate of 0% in the data analysis.

ii) To determine if the cadherin DNA of field-sampled insects is
amplified, we test for amplification of a conserved region of the
cadherin gene that occurs in all known susceptible and resistant alleles
(Morin et al. 2004).  As described in Tabashnik et al. (2006), “We
checked all pools using this approach and >99% tested positive.  Because
as few as one amplifiable allele from a pool of insects could yield a
positive result for this control reaction, we also tested a subset of
insects individually from each of the 59 field samples.  Of the 835
individuals tested, 98.6% were positive.”  

The 98.6% amplification rate of the conserved region of the cadherin
gene indicates that DNA was not amplified from 1.4% of field-sampled
insects.  We take this into account in estimating the likelihood of
non-detection by adjusting the sample size accordingly.  For example, if
1000 alleles are screened from 500 individuals and the amplification
rate of the conserved region is 98.6%, the corrected sample size is 986
(see C below).

iii) As described in Tabashnik et al. (2006): “In addition to standard
positive controls for each of the three r alleles in all tests, we
included “blind” positive controls as follows:  Two researchers
analyzed each field sample.  One researcher prepared DNA and added
individuals with one or two r alleles from laboratory-selected resistant
strains in zero to three (usually one) of the pools tested from each
field site.  The other researcher performed PCR and did not know which,
if any, of the pools contained these blind positive controls.  The
detection rate for blind positive controls was 97% (97/100).”  The
rate of false negatives (3%) caused by failure to detect r alleles
present in pools is incorporated in the estimate of the likelihood of
non-detection, as described below (C).

B. False positives.  To detect false positives, we use a standard
technique.  All tests of field samples include blanks, which are gel
lanes containing all of the PCR reagents, but no DNA.  If a blank yields
a positive result, this indicates contamination (i.e., a false
positive).  In this case, PCR reaction conditions are corrected and the
field samples are retested.  Results are included in the data analysis
only if the blanks do not yield positive results.  

Of the 5,571 field-sampled insects tested in Tabashnik et al. (2006),
none yielded positive results.  Thus, the problem of false positives is
minimal to nil.  When a pool of field-sampled insects yields a positive
result for an r allele (e.g., r2), each individual in the pool will be
tested separately to verify the positive result and to more precisely
estimate the frequency of resistance in the pool.

C. Non-detection.  As described in Tabashnik et al. (2006), the
likelihood of non-detection is estimated as follows:

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Malleles in the sample of 5,571 individuals is 0.041 = (1 – [0.0003 X
0.97] )11,142 X 0.986.”

The goal in 2006 is to screen 500 field-sampled individuals with PCR
(i.e, N=500).  Assuming no r alleles are detected and values for D and A
similar to those above, the probability (P) of non-detection is
estimated as:

i) for true r allele frequency of 0.00316 (frequency of rr = 0.00001), 

P = (1- [0.00316 X 0.97])1000 X 0.986 = 0.048

ii) for true r allele frequency of 0.01 (frequency of rr = 0.0001), 

P = (1- [0.01 X 0.97])1000 X 0.986 = 0.000067

iii) for true r allele frequency of 0.001 (frequency of rr = 0.000001), 

P = (1- [0.001 X 0.97])1000 X 0.986 = 0.38

Below please find additional discussion of the potential for
non-detection from Tabashnik et al. (2006):

“It is important to consider potential underestimation of r allele
frequency based on DNA screening.  DNA screening based solely on males
caught in pheromone traps could cause underestimation if the probability
of capture in traps was lower for rr or rs males than for ss males. 
However,  tests conducted in large cages (64 m3) in the field refuted
this hypothesis for pink bollworm (Carrière et al. 2006).  Furthermore,
DNA screening of pink bollworm from bolls, which was independent of
males caught in traps, also detected no r alleles (n = 1,344; Table 1).

If alleles other than cadherin mutants r1, r2, and r3 confer pink
bollworm resistance to Bt cotton, the results of our DNA screening could
underestimate the frequency of resistance.  For example, resistance to
Cry1Ac in some strains of diamondback moth is not linked with cadherin
(Baxter et al. 2005).  However,  in four laboratory-selected
Cry1Ac-resistant strains of pink bollworm tested so far, all resistant
individuals screened have two copies of the known r alleles (i.e., r1r1,
r2r2, r3r3, r1r2, r1r3 or r2r3) and no other resistant alleles have been
detected (Morin et al. 2003, Tabashnik et al. 2004, 2005b).  Although
the presence of additional resistance alleles at the cadherin locus or
other loci
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hic area of interest based ON spatial data

volume 1, figure 3, map 1A or 1B (electronic format)

zoom in on target area to retrieve field numbers

associated with the field(s) in question

view specific, weekly, associated field level data

from

 volume 2, table 1  [Raw data]

or

volume 2, table 2 [ave. weekly data]