If blood feeding status and duration of attachment are desirable factors for protocol guidance, effective methods have been developed to determine these parameters. Blood consumption during the early feeding phase can be verified by PCR, as well as, identification of the host blood meal source (Che et al., 2015). However, depending on the sensitivity of the assay, it is also possible to detect remnant blood from the host on which the adult tick previously fed during the nymphal stage. The duration of tick attachment and phase of blood feeding could also be scored morphologically using tick engorgement indices based on comparisons of scutal and full body dimensions = "scutal index" (Yeh et al., 1995; Falco et al., 2018). The approximate timing (in days) for initiation of blood feeding by adults feeding on mammals has been established for Ixodes pacificus/scapularis, D. variabilis, A. americanum, and R. sanguineus. Prior to and during the early blood feeding phase, when other fluids are primarily being taken up by the feeding tick, the tick cuticle does not look substantially different in color and no inflation of the body size is evident. Physically, the tick cuticle begins to change color and the tick becomes inflated during the initiation of the rapid blood consumption phase. On the day this occurs, the change in appearance may be quite rapid over the course of several hours as the "big sip" commences (body typically turns a dark gray color as it inflates).

As an example, an adult Ixodes ricinus female takes ~7-8 days to feed to repletion. The slow feeding period occurs between ~1-5 days after attachment, during which the female ingests ~1/3 of the total blood meal. Between 24-72 hours very little blood is ingested. About 2/3 of host blood is consumed during the rapid engorgement phase ("big sip"), which occurs during the last 24-48 hours before the engorged tick drops off the host (~6-8 days post-attachment) (Figure 1, Franta et al., 2010; Sojka, 2015).  PG 53
The Agency appreciates the background information. However, based on previous SAP comments, the determination of tick feeding status has been removed from the guideline because the timing of the count/removal of ticks would occur before engorgement takes place.

Public Recommendations (docket ID # in parentheses):
258
"Define what constitutes an "engorged" tick. Partially engorged, fully engorged?... "Blood-feeding to point of engorgement" is not adequately described; is it assess by scutum enlargement, increase in weight, etc.? Recommend including additional detail on "point of engorgement" to better assess criteria." (0008)
Engorgement as an endpoint for ticks has been removed from the guideline.
259
"A fact that I also disagree with in the WAAVP and EMA guidelines is discussion of engorged ticks. The study designs are setup with maximum removal time of 72h and most often within 48h. Ticks will not show engorgement within this amount of time. I have even performed squish test in my lab and could not tell the difference in body fluid results between an unfed female and a 72hr attached female R.s. tick. At most 4 categories should be counted attached and unattached in each of Live & Dead..."All references to engorged ticks should be removed. Engorgment takes 4 to 7 days and we are removing them at most 3 days after infest. You can not determine engorgement in that short time." (0032)
Engorgement as an endpoint for ticks has been removed from the guideline.
260
Re: determination of engorgement: "No, it is a subjective determination. Males do not feed to engorgement, so if you are measuring engorgement you will only have engorged females. Additionally, pathogens are transmitted well before full engorgement occurs (usually within 48 hours of attachment). In addition, ticks will not engorge in the time that they are on animals in these studies (typically only 48h) so it is not possible to accurately assess engorgement in this study design." (0026)
Engorgement as an endpoint for ticks has been removed from the guideline.
261
"Yes, blood feeding can be determined without engorgement. Squish the tick between two clear pieces of plastic and see if blood comes out. This is a similar method for what is used to determine the fed status of some biting flies. However, presence of blood does not equal engorgement for ticks. This is unlikely to be a useful endpoint for tick studies." (0026)
Engorgement as an endpoint for ticks has been removed from the guideline.

Charge Question 15
The endpoints for repellency and mortality contain numerous different categories which define how ticks should be recorded for each endpoint (e.g., repellency).
a. Are there any additional categories which might be needed for the different endpoints (e.g., repellency)? Is the separate presentation of the endpoints clear?
b. Can both repellency and mortality be assessed in a single study, or does each endpoint need to be assessed separately? Please explain.

Panel Recommendations:
262
a. The Panel did not think there is a need for more categories; rather, the Panel recommended a reduction in the number of categories. While the multiple categories offer some potential increased resolution into the responses of the ticks to product testing, the number of categories would be unrealistic in practice.  PG 54
The Agency agrees with this comment and the number of categories has been reduced.
263
Efficacy Endpoints
The proposed guidelines state: "Mortality should be determined by comparing dead, unengorged ticks on the animals and in trays/cages to the control group. Ticks that feed to engorgement can transmit agents that may cause disease; therefore, mortality after blood-feeding to the point of engorgement should not be considered a viable endpoint."

The Panel suggested revising this to indicate that mortality at or before 48 hours would be a reasonable endpoint for mortality to prevent pathogen transmission, if that is the goal.  PG 55
The guideline has been revised to incorporate this comment. Since tick counts for mortality testing are to be conducted by 48 hours, which is before "blood-feeding to the point of engorgement" would happen (or possibly any blood-feeding for that matter), the endpoint for the tick mortality testing has been changed to "live ticks on the treated animal vs. live ticks on the control animal."
264
Efficacy Endpoints
The proposed guidelines state: "For testing repellency against ticks, a repellent effect is a reaction by a tick to avoid contact with a treated animal. Since repelled ticks will move away from the treated animal and possibly be unrecovered, repellency should be determined indirectly by counting the number of live or dead ticks (attached or unattached) on the animal plus live or dead ticks in the cage that are engorged compared to the control animals. Moribund ticks should be considered alive. Other methods of evaluating repellency against ticks (e.g., hot foot, inhibition of attachment, disruption of attachment; Halos et al. 2012) may also be considered but should be justified."

The Panel noted that the time frame is important here. If following EMEA guidelines (European Medicines Agency, 2016), the ticks should not attach to the host within a 24- hour time frame. Therefore, host and cage surroundings should be checked for ticks and numbers on and off the host need to be compared to control animals. Off-host live, or dead ticks could both be counted. Antifeedant could be considered as an additional endpoint. The number of live and unattached ticks would be a measurement of this.  PG 55
The methods for tick repellency have been revised to have the ticks placed near the animal instead of directly on the animals and will compare the number of ticks (dead or alive) on the animal 24 hour later to determine repellency.
265
Data collection and recording for ticks
For mortality trials, the mode of action of the product would dictate when the data is collected. Systemic products must be ingested by ticks for mortality to occur and could take several days to act based on the prolonged feeding period of ixodid ticks. The Panel noted that a time point of 48- 72 hours should be reasonable to capture tick mortality with systemic acaricides. For some acaricides that act much sooner, earlier measurements should be made. Comb counts should be specified to include tick removal, unless mortality measurements are being made for retained ticks over longer time points in studies.  PG 55
For mortality trials, the 48 to72 hour counts will remain, with the option to count sooner. The guideline has also been revised to clarify tick removal as suggested.
266
Data collection and recording for ticks
For the tick mortality effect categories, as the Panel has noted previously, all instances of "engorged" should be removed and replaced with "partially fed." The Panel observed that it seems like there are too many categories here to be truly useful. Reducing the number of categories to evaluate would simplify study interpretations and perhaps reduce the sample sizes of ticks and animals needing to be used. The simpler categories from the European Medicines Agency (EMA, 2016) would probably suffice for most study goals. Also, it seems questionable for moribund ticks to be counted as alive; it is probably more accurate to count them as dead because it is unlikely that they will adequately recover to the point of being capable of re-attaching to a host. Holding moribund ticks off the host for at least 24 or up to 48 hours, under optimal temperature and humidity conditions for survival would allow the study to differentiate live from dead ticks unambiguously. For the tick repellency categories, the Panel agreed the data should typically be collected within 24 hours post-infestation, though sometimes ticks may take longer to attach to the host; up to 48 hours may be appropriate to evaluate repellency in some host-tick systems. Also, there are too many categories for tick scoring here. Within 24 hours, there is no blood acquisition by ticks, and so these categories are not accurate. Blood feeding as a factor for repellency should be removed when using short time frames.  PG 55-56
The number of categories in both mortality and repellency tick testing has been reduced. Since the final counts for the mortality endpoint are already at 48 hours, "moribund" ticks will still be counted as "live."

Since the repellency endpoint will now be whether the tick is on or off a host animal at 24 hours, the blood feeding category will be removed.
267
Data collection and recording for ticks
In mortality testing studies, how can host immune response to ticks be differentiated from the effects of the product tested? The Panel noted that it may be important to provide guidance for individual variations that occur in host rejection of ticks due to an immune response vs. product induced mortality determinations during trials as an endpoint (European Medicines Agency, 2016; Wada et al. 2010). This may not be an issue for R. sanguineus, but it could be the case for other tick species tested, such as I. scapularis (Szabo and Bechara, 1999; Gebbia et al., 1995).  It is possible to have large numbers of ticks attach to a host, over the 25% threshold, and rejection by the host may occur during feeding trials that is due to host immune response, not due to the product's effects. Monitoring of host inflammation around feeding ticks would provide insight regarding product-induced mortality vs. immune-induced mortality. Animals with noted inflammation around bite sites should be removed from the study to prevent confounding of results. This could be of particular importance as animals are infested multiple times over the course of several months and may develop acquired immunity to tick feeding during later trials.  PG 56
While this comment is appreciated, it is outside the scope of the guideline to differentiate host immune responses.
268
Data collection and recording for ticks
b. Optimally, mortality and repellency should be determined in different studies with different designs, because they are different outcomes and have the strong potential to bias the outcome. The current methods muddle the distinction between mortality and repellency. For example, if 50 ticks are put on an animal, and at 24 hours investigators find 10 dead ticks on the animal, 0 live ticks on the animal, and 40 moribund ticks in the cage, this could be scored as a repellent, when equally likely this was a slow acting acaricide without repellent activity. While some materials are both toxic and can act as a repellent, other materials may do one or the other. Thus, the Panel suggested that the design for repellency assays needs reconsideration, as discussed in more detail in the response to Charge Question #10.  PG 56
These guideline sections have been revised. Based on the revised methods, assessing repellency would likely not be able to be combined with mortality.
269
Data collection and recording for ticks
b. The Panel also recommended that for a repellency study, ticks should not be placed directly onto an animal. Instead, ticks should be placed in the local environment with the animal to allow ticks to naturally attach to the host; if there is a repellent effect, fewer ticks will be found on the animal (Dumont et al., 2015b).  PG 56
The guideline has been revised to incorporate this comment.

Public Recommendations (docket ID # in parentheses):
270
"Adult mortality advises live fleas vs. dead ticks to determine mortality. Recommend assessing by counting the live fleas only (not moribund) and comparing treated vs. untreated groups. Additionally, this should be for all "kills" claims not just "starts killing". (0008)
See response to comment #14.
271
"Calculation of Repellency differs EMA guidelines and the proposed formula includes dead ticks in the control & treatment retention rate Efficacy = (Attached ticks on control - Attached ticks on treated)/(attached ticks on control) x 100" (0008)
The guidelines have been revised to be consistent with the EMA guidelines.
272
"Tick efficacy should be based on live ticks on dogs, similar to flea efficacy. With that approach, there is no need for collecting ticks that fall off the animals as this is inherently variable with ticks escaping and/or being lost in feces, urine, vomitus, etc... Can the EPA justify why tick efficacy would be based on dead ticks? Tick efficacy is routinely calculated based on a comparison of live ticks on the treated dogs compared to the control dogs (similar to flea efficacy as described in this draft guidance). Quantifying dead ticks is highly variable and inaccurate. Measuring live ticks is much more accurate and appropriate for evaluating tick efficacy." (0026)
See response to comment #14.
273
"Yes, repelling ticks is a viable endpoint. Companies need to be given a chance to develop products and formulations that give the end user a variety of options in order to control tick populations. b. This would have to be a defined endpoint in the protocol before trial initiation. Methods to assess repellency can be very similar to mortality assessments. c. Yes, it can be. The initial study question must be defined in the protocol before the initiation of the study. As the methods and endpoints are similar ratings for repellency versus mortality then these need to be tested by two separate trials i.e., one trial to determine mortality effects and one trial to determine repellency effects." (0008)
See response to comment #268.
274
"The second edition of the WAAVP guidelines (published in 2013) updated the tick categories to only four categories: live free, live attached, dead free, dead attached. Engorgement status was removed from consideration as most ticks will not be engorged during the time that they are on dogs in laboratory studies and as it can be difficult to assess when ticks are not fully engorged. Suggest that the EPA guidelines should be aligned with this to ensure all data collected is meaningful and adds value (in regard to engorgement status) and to allow studies to be used in multiple geographies." (0026)
We agree with this comment and these sections have been revised to align with the EMA guidelines.
275
"Based on the current endpoints. It seems that the mortality endpoint is dead ticks compared to the control animals, where repellency is looking at live and dead ticks, so they can be assessed in the same study. If counts are done at 24-72 hours post infest or treatment, there shouldn't be engorged ticks in kennels." (0026)
See response to comment #268.

METHODS FOR TESTING AGAINST MOSQUITOS OR BITING FLIES (SECTION (L))

Charge Question 16
Comment on the adequacy of outcrossing lab-reared colonies of mosquitoes every 3 years.
a. Is three years an adequate timeframe to retain genetic diversity in lab-reared populations? Why or why not?
b. If not, how often should lab-populations be outcrossed with wild-type mosquitoes? Please provide references if relevant.

Panel Recommendations:
276
What is the goal of periodic outcrossing of mosquito (as well as other ectoparasite) colonies? The Panel observed that retaining genetic diversity is irrelevant for toxicology testing (unless for example, examining frequency of resistance alleles) and questionable for behavioral studies. Most insect species including mosquitoes do not die off with inbreeding. Mosquito colonies at the USDA laboratory in Gainesville, FL have never been outcrossed and have been used successfully for a variety of tests for years. If the goal is to maintain some part of a large population with relevant proportions of resistance alleles, then the Panel found that this protocol is not detailed enough.  A recent paper by Ross et al. (2019) concludes that inbreeding depression occurs in this species due to low population size. Colonized mosquito populations suffer inbreeding depression and adaption to laboratory conditions. Laboratory environments are inherently artificial, and colonized mosquito populations experience an entirely different set of selective pressures compared to natural populations, which could lead to declines in reproduction (Bryant and Reed.,1999). Ross et al. (2017) suggested that while life history traits of Ae. aegypti do not change consistently with laboratory maintenance, traits where selective pressures are absent in the laboratory, such as flight ability, feeding behavior, and thermal tolerance might still be compromised.

Ross et al. (2019) concluded that variation in fitness and performance was dependent on colony size, but that most individual traits were unaffected, and patterns of adaptation were not consistent across populations. Performance of laboratory populations of Ae. aegypti maintained in a larger population increases. Rates of laboratory adaptation can be slowed or minimized by using more natural rearing environments such as maintaining large populations (400 vs. 100 in larger cages at a variable temperature and environments that are more complex.

The Panel agreed that many traits are lost in colonized insects shortly after they are selected for colony conditions. However, genetic conditions tend to stabilize. Adding new (uncharacterized) genetic material periodically would change the genetics of the colony so they are different than they were when the colony was used for previous studies. This could produce different results in certain types of tests.  PG 57
The outcrossing recommendation has been removed.
277
Rather than periodic outcrossing, the Panel suggested testing a standard strain that is well-studied and has a history in the scientific literature and comparing the counts of live mosquitoes on treated versus untreated animals. Believing that there will be widespread differences between strains assumes facts not in evidence. Furthermore, every time a strain is outcrossed the strain changes and could result in non-reproducible results, a major drawback to any manipulations of the strain used for testing. In addition, outcrossing with uncharacterized field-collected individuals could introduce pathogens, as well as, resistance to ectoparasiticidal compounds.  PG 58
The outcrossing recommendation has been removed.
278
If the Agency decides to include protocols regarding outbreeding in the updated guidelines, the Panel cautioned that it would be more appropriate to suggest outbreeding after every so many generations than to specify a time period (i.e., every 3 years). Several articles indicated that the simplest way to maintain the fitness of colonized mosquito populations, especially Ae. aegypti, is to cross laboratory colonies to an outbred population. Yeap et al. (2011) suggest a 2-year period for out-crossing, but a rationale was not readily discernable from the article.

With an estimated generation time of approximately 30 days for Ae. aegypti, (Sowilem et al.,
2013), a 3-year outcrossing interval could result in >36 inbreed generations.  However, Ross et al. (2019) established replicate populations of Ae. aegypti mosquitoes and maintained them in the laboratory for twelve generations at different census sizes [<100 individuals or 400 individuals]. At F5, life history traits were compared between replicated large populations and inbred lines. After two generations of brother-sister mating, the inbred lines had reduced fitness relative to the large populations, with substantial costs to larval survival and development time. Although the study design involved a comparison among replicates established from eggs from an ancestral population, the data suggest adaptive changes to laboratory conditions in the F5 generation. Therefore, if outbreeding was important, it should be done more frequently than on a 3-year basis.

Alternatively, Ross et al. (2017) describe a simple protocol for maintaining Ae. aegypti mosquitoes in the laboratory, which minimizes laboratory adaptation and implement outcrossing to increase the relevance of experiments to field mosquitoes. They suggest outcrossing be repeated for at least three consecutive generations to produce colonies that are at least 87.5% similar, genetically, to the field population.  PG 58
The outcrossing recommendation has been removed.

Public Recommendations (docket ID # in parentheses):
279
"Outcrossing laboratory colonies with field populations every 3 years is not appropriate. The idea behind having an established laboratory colony is to have a "reference" colony to which things can be compared back to over long periods of time. The outcrossing of field populations has the potential to introduce pathogens which would deem the colony unacceptable for use. Additionally, it poses the risk of introducing an insect pathogen that could destroy the entire colony. If claims to control resistant populations are sought, separate tests should be conducted on arthropods with known resistance to the active ingredient (or mode of action) to support such claims. If the intent is to introduce "alternate" genes into the colony, these alternates will be diluted in a short period of time due to the generational efficiency of most mosquito species." (0008)
The outcrossing recommendation has been removed.

Charge Question 17
In the interest of reducing vertebrate animal testing, we propose simultaneous testing of up to three mosquito species on a single vertebrate animal.
a. Is simultaneous testing of three mosquito species feasible? Why or why not?
b. If simultaneous testing of three mosquito species on a single vertebrate animal is feasible, then please comment if the proposed methods are adequate, and if there are alternative methods please provide detailed methods and references.
c. If testing three species at once is not feasible, provide additional detailed methods and associated references to limit the number of vertebrate test animals necessary to complete efficacy testing against the three required mosquito species and maintain adequate statistical power.

Panel Recommendations:
280
After consulting mosquito experts (e.g., Drs. Dan Kline and Bill Reisen, personal communication), the Panel determined that simultaneous testing with three mosquito species should not affect the test results. The mosquitoes do not compete for feeding sites and do not otherwise interact in ways to alter results expected with a single species. Proposed methods look reasonable, but it would be nice to reduce the number of recommended vertebrate animals used.  PG 59
No changes to the guideline are necessary.
281
After an extensive discussion of this Charge Question, the Panel recommended the following: If three mosquito species are tested simultaneously, 50 mosquitoes of each species should be used. .......  PG 60
No changes to the guideline are necessary.
282
After an extensive discussion of this Charge Question, the Panel recommended the following: ......... For flies known to exhibit aggressive behavior and (or) inflict a painful bite (e.g., stable flies), efficacy testing conducted with 50-100 flies may not be recommended because of the adverse effects on the vertebrate animals involved. Under natural conditions, stable flies rarely complete a blood meal on the first try because they are dislodged by the host because the bite is painful. The fly merely flies around the host and tries again. This continues until the fly is replete. When the fly is undisturbed, as it would be under laboratory testing conditions, the time to repletion averages 147 seconds, as documented Schofield and Torr (2002).  PG 60
The models were re-run for stable flies using 25 flies and determined if 25 flies per animal is desired then 12 dogs per study should be used. The infestation time has also been revised to only for 30 minutes.

Public Recommendations (docket ID # in parentheses):
283
"For a general mosquito claim, product must be effective against 3 species of mosquitoes, some products may repel, and others efficacy is measured by landing, protocol discusses testing multiple species with single test. We do not believe this would be possible." (0008)
The mosquito section allows the testing of multiple species at the same time; however, the study does not have to. No changes to the guideline are necessary.

Charge Question 18
The proposed methods to test mosquitoes and biting flies indicate vertebrate test animals should be sedated prior to exposure to invertebrate pests.
a. In the interest of minimizing stress of sedation on vertebrate test animals please discuss whether sedation is necessary. If sedation is not necessary, please provide detailed methods and references and discuss the strengths and weaknesses of the methods for conducting efficacy testing.
b. If sedation is necessary, please provide methods to minimize the number of times vertebrate animals should be sedated.

Panel Recommendations:
284
Because the efficacy of a product is being evaluated, a sedated animal allows this evaluation to be conducted without the effects of host defensive behaviors. An un-sedated animal confined in a 2 ft3 cage with 50-100 flies or mosquitoes feeding on it would be in constant motion. Pain associated with the bites of mosquitoes and stable flies would interfere with an accurate evaluation of product efficacy and the pain would be difficult for a test vertebrate (e.g., a dog) to endure for 1 hour. The Panel suggested reducing the number of fly exposure trials requiring animal sedation to every 2 weeks (Days 7, 28, etc.), when possible.  PG 60
It is important to balance realistic testing with humane treatment to the host animals to produce reliable results. The proposed guideline already reduced infestations for mosquitoes and flies compared with fleas. Currently, for a typical 1 month product, there are three infestations: day 3, day 14 and day 28. This will provide enough endpoints to gain reliable data with the reduction of 1 infestation point. No changes to the guideline are necessary.
285
If starved insects are used, this should cause more rapid feeding on the test vertebrate, and sedation time and exposure to the insects might thus be reduced. Young (3 to 5 days old) stable flies are routinely starved for 4 hours before a test where feeding is involved. Stable flies will remain alive for 24 hours or more if given water on cotton ball or pads (Jones et al., 1992). Stable flies will live several days if provided with a 10% sucrose solution or Gatorade (powder is easy to keep and store). Mosquitoes are not usually starved but maintained on water or 10% sucrose.  PG 60-61
See response to comment #18.
286
If one of the criteria for a successful test is the presence of blood in the insects' gut, the Panel observed that small amounts of blood would be sufficient. Insects do not need to feed to repletion. Stable flies can fill up in an average of 147 seconds (Schofield and Torr, 2002). Some mosquitoes can fill up in about <= 3 minutes, if undisturbed.  PG 61
No changes were proposed in this comment.

Public Recommendations (docket ID # in parentheses):
287
"Selection and allocation of test animals. Methods other than sedation of animals should be considered. Sedation can be hard on the animals, especially if there are multiple data points over long periods such as a collar evaluation." (0008)
Sedation of animals is only suggested when testing against mosquitoes as they are difficult to count on moving animals and against painful/aggressive biting flies. These pest claims are rarely proposed. Sedation should not be used for testing against other pests. No changes to the guideline are necessary.
288
"The effects of sedatives can differ -- some pain medications have sedative properties, but not all sedatives have pain management properties. In addition, not all sedatives are anxiolytics, meaning that some sedatives prevent movement but mental anxiety is still present. If the guidelines recommend to sedate animals, they should also provide that the animals are administered pain medications or a drug that has proven analgesic and sedative properties." (0019)
The following sentence has been added:

"For infestations using painful or aggressively biting mosquitoes or biting flies, the animals should be administered pain medications or a drug that has proven analgesic and sedative properties."
289
"Sedation is considered necessary in this model to prevent animals from interfering with the feeding of the mosquitoes or flies." (0026)
 This feedback has been incorporated into the guideline. See response to comment #288.

Charge Question 19
Comment on the timing of exposing vertebrate test animals to mosquitoes and biting flies.
a. Are the number of exposures for products with different durations of efficacy adequate for determining efficacy? Why or why not?
b. If the number of exposures to pests can be decreased, please indicate specifically which exposures can be skipped. If efficacy should be evaluated at more time points, indicate when exposures should occur and for what product types (e.g., spot-ons, collars, and residual shampoos), and discuss the value provided by the additional time points in relation to the additional stress on test animals from additional sedations.

Panel Recommendations:
290
a. The proposed guidelines variably specify that each test vertebrate animal should be exposed to 50-150 adult, pathogen-free mosquitoes or biting flies for one or more time points. As has been previously mentioned, the Panel questioned whether adequate numbers of all insect species listed in the proposed guidelines are readily available for this.  PG 61
Language has been added to indicate that flies may be obtained from wild populations if they are not readily available in lab colonies. In addition, the specification of "pathogen-free" has been revised to "free of vector-borne pathogens" to be more specific.
291
a. Furthermore, the Panel strongly recommended that the requirement for exposing vertebrate test animals to 50 stable flies be rewritten due to the extreme pain and distress caused by exposure to this aggressive biting species. Even decreasing the number of stable flies from 50 to 25 may still be excessive for an individual animal to tolerate for the proposed one-hour exposure period, let alone multiple exposures. The reduction in power and corresponding need for an increased number of vertebrate test animals caused by decreasing the pest number to 25 stable flies, and potentially even lower, should be further considered through modeling. The Panel was interested in seeing just how much decreasing the number of stable flies would increase the number of necessary test animals. Similarly, the number of control animals should be related to the expected degree of response. Since the control mortality would be expected to be very low in the case of stable flies, the Panel suggested that perhaps fewer control vertebrate animals could be used.  PG 61
The stable fly model was re-run using 25 stable flies vs. 50 and the guideline has been revised to add that if 25 flies are tested vs. 50 then 12 dogs per groups should be used. The guideline has also been revised to state that the exposure time for flies should be 30 minutes rather than 1 hour. The guideline already had reduced amounts of exposures (only days 3, 14 & 28.) Finally, language with sedative and pain medication suggestions to help alleviate pain and stress to the animals has been added.

292
a. As previously raised by the Panel in discussing other charge questions, better definitions of "adequate infestation" of test animals, preferably specific to both pest and test animal species, would be another strategy for reducing overall sample sizes. The Panel also discussed specifying appropriate expected levels of blood feeding on control animals. The figure of 75% currently appears in the "Sample Size for Pet Product Studies" document (page 7) and the Panel found that it was reasonable to expect that this would be achieved quite quickly. This would be especially true when using flies in their most active 3-5 days of age period and after adequately starving them for 3-4 hours prior to exposing them to the test animal. In both cases, better-refined definitions and study design recommendations may also increase the power of the resulting data and allow further reduction of vertebrate animal numbers.  PG 61-62
The guideline currently includes some parameters to assist in increasing the power of the data, such as requiring a minimum amount of bloodfeeding or retention of arthropod pests on control animals, which were incorporated into the power versus sample size modeling.
293
b. If the registrant can reasonably expect that their product will last a full month (as a repellent and/or with insecticidal activity), the Panel found that it may be permissible to allow testing to occur at Day 7 to show the topically applied product has spread over the dog and then eliminate Day 14 exposures. Day 21 and Day 28 exposures should still be included. The Panel noted that this eliminates one time point (Day 14) and still proves month-long residual efficacy.  PG 62
For products lasting 1 month or less, all the time points listed in the guideline will remain because collection at multiple time points helps with the interpretation of a dataset containing the occasional efficacy failure. For a mosquito or biting fly product that is expected to last 30 days, testing had already been omitted on day 7. However, it should be noted that reducing the number of infestations does not reduce the number of animals used nor the overall duration in each study.
294
b. The Panel recommended that the guidelines address the duration of each experimental exposure. Many of the Panel's concerns about animal welfare related to both sedation and exposure to biting flies could be addressed by limiting the exposure time. A 30-minute to one-hour exposure time for stable flies was suggested.   .............  PG 62

b. The Panel recommended that the guidelines address the duration of each experimental exposure. Many of the Panel's concerns about animal welfare related to both sedation and exposure to biting flies could be addressed by limiting the exposure time. A 30-minute to one-hour exposure time .......... The same exposure times should also be applied to the enrollment phase of these studies.  PG 62
See response to comment #291.

Public Recommendations (docket ID # in parentheses): n/a

Charge Question 20
Comment on the methods for assessing mosquito and biting fly blood-feeding.
a. Are the proposed methods adequate to determine blood feeding for insects where blood feeding status cannot be visually observed through the abdomen?
b. Are there other methods for determining if insects blood-feed? If so, please provide additional methods and references.

Panel Recommendations:
295
a. The Charge Question seems to be asking if sampling half the live flies (where blood feeding cannot be determined by observation) with a blood smash test is acceptable.
This question is subjective and depends on what level of risk is acceptable, particularly where transmission of pathogens is of concern. The remainder of the flies which are to be checked at 48 hours post treatment could be evaluated for blood using one of the more sensitive blood detecting techniques described below, at which time all the flies would be evaluated, thus eliminating uncertainty.  PG 62
The guideline has been revised to determine blood feeding for all mosquitoes (for mortality and repellency claims).

In regards to using more sensitive techniques, the following language has been added:  "It is preferred that blood-feeding is assessed and confirmed for each individual insect using an enzyme-linked immunosorbent assay (ELISA) (e.g., Beier et al. 1988) or polymerase chain reaction (PCR) based method (e.g., Fitzpatrick et al 2019); however, other methods such as squishing may also be appropriate as long as the method chosen is used consistently throughout the study and applied the same to treated and control groups."
296
Of concern to the Panel was that EPA's definition of "repellency" (has not imbibed blood) differs from the traditional entomological definition of repellency which is directed movement away from repellent. The proposed guidelines (i.e., on page 18) seem to confuse repellency and feeding inhibition. The guideline states:

"i. Repellent effect. Insects should be evaluated to determine if they took a blood meal. Insects that take a blood meal are not repelled. More conservative endpoints such as insect landing may also be considered but should be justified."

"i. Repellency. To determine repellency only, immediately after removing animals from the cage all insects should be aspirated from the cage, knocked down (e.g., using CO2, or frozen), identified to species, crushed on a light background to assess for blood-feeding."

The Panel noted that this does not conform to definitions of repellency put forward by several groups. Repellents cause oriented movements away from the source. In relation to feeding, a better term might be a feeding inhibitor (Deletre et al., 2016). For example, the European guidelines define repellent effect as "a product with a repellent effect will cause the parasite to avoid contact with a treated animal completely and/or to leave a host" (EMEA/CVMP/EWP/005/2000-Rev. 3).

Marchiondo et al. (2013) writes, "Two types of repellency are defined: "sensu stricto" for repellency characterized by an irritant effect, causing the tick to move away from the treated animal or leading it to fall off soon after contact with the treated hair coat within ∼6 - 8 hours and "sensu lato" for all other tick repellency (or expellency) up to ∼24 hours. The first, repellency sensu stricto, may be attributed to the vapor phase of a compound or irritant effect through direct contact (e.g., oil of Citronella but also some synthetic pyrethroids), while the second repellency (or expellency) causes inhibition of attachment or detachment of already attached ticks, (e.g., some synthetic pyrethroids or amitraz)."

Collection of unfed flies means either: 1) the flies simply didn't feed during the test period, 2) the test material was a true repellent, or 3) the test material was a feeding inhibitor.  PG 62-63
We agree with the panel's definition of repellency and have clarified the definition vs. the appropriate end point to test the product. The guideline recommends that blood feeding for mosquitoes and biting flies should be the endpoint since it is easier to demonstrate. However, landings as the endpoint is also an option, if justified. As for fleas, the referenced literature was considered, and a statement was added to the guideline to encourage the submission of a protocol for flea repellency tests.

297
b. The Panel noted that techniques used to determine blood feeding in cat fleas include Drabkin's technique to determine the quantity of blood consumed by adult cat fleas (Rust et al., 2002) and should be applicable to flies. Several other procedures of determining blood are outlined in Marchiondo et al. (2013). A real-time PCR analysis of blood feeding in fleas was 10,000-fold more sensitive than the Drabkin technique (Wang et al., 2012). Wang et al. (2012) concludes, "The HMBS PCR method developed here offers the advantages of both exquisite sensitivity and specificity that make it superior to other approaches for quantification of blood ingested by fleas. The capability to detect minute quantities of blood in single fleas, particularly immediately after colonization of the host, will provide a superior tool for studying flea-host interactions, flea-borne disease transmission, and flea control strategies."  PG 62-63
See response to comment #295.

Public Recommendations (docket ID # in parentheses):
298
"In section l(5)(ii)(a) I dont see how the abdomen of live mosquitos can be visually assessed and then the mosquitos held for the additional 48 hours post exposure. When they are flying and moving, they cannot be properly assessed in this manner. The only proper way to do this is killing then by freezing and then squashing them to determine blood meal." (0012)
The guideline has been revised to assess all recovered mosquitoes for blood feeding.

Charge Question 21
To reduce the amount of vertebrate animal testing, we propose to allow assessment of repellency and mortality within the same study by evaluating half of the insects for repellency before determining mortality.
a. Discuss the viability of evaluating both endpoints in the same study by using a subset of the total mosquitoes to determine repellency and provide any additional methods that would determine both endpoints within the same study.
b. The proposed endpoints for mortality are secondary to repellency (i.e., for mortality to be assessed repellency should be observed first) because a mosquito or fly that is not repelled may blood-feed and could potentially transmit pathogens that cause disease. Discuss the potential value of assessing mosquito mortality that are not also repelled. Should mortality of mosquitoes that blood-feed be considered a viable endpoint to show efficacy of a repellent-product? Why or why not? If yes, provide specific rationale as to why.

Panel Recommendations:
299
a. The Panel observed that if the timing of endpoint determinations is different, it seems reasonable to study both repellency and insecticidal activity against mosquitoes and biting flies during the same test. Repellency is immediate; knockdown/mortality occurs up to 48 h after exposure. One is, however, looking to see that none of the insects had a blood meal if the products are to be effective in either category.

As stated in the proposed guidelines (OCSPP 810.3300) Section (l) (4), efficacy endpoints, the "endpoints (e.g., blood feeding, mortality) should be selected with respect to the intended use pattern for the product and mode of action of the a.i."

An active ingredient (a.i.) could have only: (a) a repellent effect; (b) a repellent and a toxic effect (e.g., permethrin); or only a (c) cidal effect (e.g., fipronil).  Thus, the a.i.'s mode of action and its intended use pattern should drive the testing methodology and the efficacy endpoints.

The goal of insecticides/repellents applied topically to dogs/cats is to protect the animal from the biting adult stages of mosquitoes and biting flies and to interrupt blood feeding and thus significantly reduce the potential for disease transmission. Assessment of blood feeding is the critical thing to assess in live, knocked down, moribund and or dead insects after being exposed to a topically treated dog/cat.  Thus, if based on blood feeding >=90% of the insect species being assessed are alive and have no blood in them, then this a.i. has a mode of action that should be considered a repellent. If the a.i. has both a repellent and a mortality effect, then >=90% of the recovered insect species whether they are alive, dead, or moribund should have no blood in them since the a.i. either repelled them and/or killed them before a blood meal could be taken.

Therefore, the Panel concluded that looking at subsets (e.g.-removing half of the insects) of the total number of mosquitoes and biting flies and looking for repellency before assessing mortality is not needed. Blood feeding should be assessed in all recovered insects and this should drive the label claim (use pattern). Depending on the extent of blood feeding the product can claim to be a repellent or an insecticide/repellent or a standalone insecticide that works so fast that insects that land on a treated dog/cat are knocked down or killed so quickly that they can't obtain a blood meal before dying or becoming moribund and subsequently dying.  PG 64
Both repellency and mortality can be assessed in the same study and the guideline has been revised to have all mosquitoes or biting flies assessed for blood feeding.
300
a. The Panel suggested that some of the automated methods be used to detect the presence of blood in an insect to confirm if blood feeding has occurred. For example, confirmation of a consumed blood meal could be performed using an enzyme-linked immunosorbent assay (ELISA) following the procedure described by Beier et al. (1988).  There are likely other references that could be cited for these methods as well.  PG 64
EPA agrees with the suggestion and has included language indicating that these automated methods including the ELISA method provided are acceptable options to detect blood-feeding.
301
b. The Panel noted that blood feeding should not occur if the test product is effective either as a repellant or as a knockdown/mortality agent. As noted in the Charge Question, repellency is primary and needs to be observed before mortality is determined. However, knowing if insects die quickly could be important. If the biting insect is not repelled and blood feeding occurs, there is potential for disease transmission unless killed quickly. Therefore, even if mortality occurs after insects are on the test animals, mortality/knockdown could also be considered as a viable endpoint.  PG 64-65
EPA agrees that absent blood-feeding, mortality is a viable endpoint and as such, will allow for assessment of mortality if blood-feeding does not occur.
302
The Panel did not agree with the assumption that the proposed endpoints for mortality are secondary to repellency. As stated above, the mode of action of the a.i. is going to define the endpoint (and label use pattern) and the ultimate goal of the a.i. is to significantly reduce or totally inhibit any blood feeding by mosquitoes/biting flies exposed to a treated dog/cat. In this Charge Question, the Agency states that "a mosquito or fly that is not repelled may blood-feed and could potentially transmit pathogens that cause disease."  This may be true but if the a.i. has a predominant killing or mortality effect, the insect can be killed or incapacitated so quickly after landing on the host that blood feeding doesn't occur.  PG 65
The guideline allows for assessment of mortality if blood-feeding did not occur.  Although mortality may not be secondary in an instance where a killing effect occurs so fast that the insect does not blood feed, the evaluation of this under the guideline study would be the same as for a repellent product that also has insecticidal claims. No changes to the guideline are necessary.
303
To respond to the first bullet point in Charge Question (b),the Panel would again refer the EPA reviewers to the previous discussion points that blood feeding should be assessed in all recovered insects and this should drive the label claim (use pattern) as a repellant, and/or an insecticide/repellent or a standalone insecticide that works so fast that insects that land on a treated dog/cat are knocked down or killed so quickly that they can't obtain a blood meal before dying or becoming moribund and subsequently dying.  PG 65
The Agency agrees and has revised the mosquito/biting fly section to provide a method to evaluate all insects for bloodfeeding and mortality in a single study.
304
To respond to the second bullet part of Charge Question (b), the Panel concluded this question would only apply to an a.i. with a mode of action that has both repellency and a mortality (toxic) effects. Under this scenario, if, based on blood feeding, >= 90% of the insect species being assessed are alive and/or dead (demonstrated mortality) and have no blood in them, then this a.i. has a dual mode of action (repellent combined with mortality effects, such as permethrin). Thus, if <= 10% of live/dead insects do have blood detected in them, then the 90% of insects that don't have blood detected will be making a significant impact on reducing disease transmission and reducing irritation due to the biting process by the insects and potential immune reactions at the bite site. No product in a real-world biological system will consistently provide 100% efficacy, thus, the Panel observed that having a minimum threshold of >= 90% seems appropriate.  PG 65
EPA agrees with the 90% threshold for this type of study.
305
Additional Comments related to Assessing Efficacy in Mosquitoes and Biting Flies
One committee member suggested that the EPA should consider adding a new section to this part of the revised guidelines that assesses knockdown/mortality on topically treated dogs/cats, regardless of their blood feeding status.  This would be analogous to efficacy assessments performed on treated dogs/cats infested with fleas and ticks where they ingest blood prior to being killed by the topically applied a.i. The primary concept is to kill these insects in their local environment which negatively affects reproduction and helps to interrupt the life cycle of the pest and reduces the overall population of the insects in the pet's local environment and also potentially reduces human exposure to bites. It has been documented in the scientific literature that some mosquitoes do not move significant distances from where they breed, thus killing local populations of these insects can be just as important as preventing blood feeding. Since only small numbers of biting insects are actually carrying infectious agents, the impact of significantly reducing blood feeding and reducing transmission of infectious agents is minimal.  PG 65
These products are not intended for controlling mosquito populations in a local area.  Additionally, EPA is not aware of, nor has evidence been provided, that these products have measurable effects on local mosquito populations or offer effective protection against mosquitoes to other animals which did not receive a dose within the local area. For these flying pests, efficacy is based on protection of the animal to which the products are applied.

Public Recommendations (docket ID # in parentheses):
306
"Yes, repelling mosquitoes is a viable endpoint. Companies need to be given a chance to develop products and formulations that give the end user a plethora of options in order to control mosquito populations. b. This endpoint would have to be defined endpoint in the protocol before trial initiation. Methods to assess repellency can be very similar to mortality assessments. c. Yes, it can be. The initial study question must be defined in the protocol before the initiation of the study. As the methods and endpoints are similar ratings for repellency versus mortality then these need to be tested by two separate trials i.e., one trial to determine mortality effects and one trial to determine repellency effects." (0008)
EPA agrees that repellency is a viable endpoint. EPA has taken the advice of the SAP in that both repellency and mortality can be assessed in the same study by assessing blood feeding for all the mosquitoes or flies.

Charge Question 22
Efficacy testing is based on the specific label claims proposed for product marketing. Common claims are "kills," "repels," and "controls" (i.e., residual efficacy). For most pesticide products (e.g., direct sprays and residual applications) the typical endpoint to support claims of "kills" or "controls" is based on dead arthropods. To evaluate dead arthropods for shampoo products we propose pests should be shampooed in a tub with a screen over the drain to keep fleas and/or ticks in the tub. This may differentiate the pesticidal effect from the effect of pest removal, which could also occur by using a non-insecticidal shampoo. Discuss the value of using counts of dead arthropods vs. using counts of live arthropods on treated compared to untreated vertebrate animals to determine different endpoints (e.g., kills).
a. Is assessing live arthropods on treated vs. untreated animals an adequate and practical endpoint for determining mortality? Why or why not?
b. Are there alternative methods for determining mortality for shampoo products? If so please provide the methods and supporting references.

Panel Recommendations:
307
a. The Panel noted that the efficacy or effectiveness of treatment is appropriately determined by comparing the proportion of pests killed by the treatment (i.e., proportion of living in the control group - proportion living in the treatment group) with the proportion of pests living in the control group (Abbott, 1925). Determining the proportion of live pests in the control group, and the proportion of live pests in the treatment group, requires counts of living pests on each vertebrate animal and knowledge of the total number of pests applied to each vertebrate animal. Ideally, all the arthropods applied to the vertebrate animals would be counted and classified as live or dead to assess the proportion of live arthropods in the control or treated group. Counts of the dead arthropods would support the claim of killing for the tested product. However, it is unlikely that the total number of arthropod pests applied to each vertebrate animal will be recovered during the counting process. Fleas originally applied to a cat or dog may be ingested or hide within or upon a vertebrate animal. Accordingly, it has become standard practice to use Abbott's formula with live arthropod counts.

In the proposed guidelines, approximately the same total number of arthropods is applied to each animal. It is reasonable to assume that the total number of pests on the vertebrate animal does not change appreciably between the time of application and the time of counting. If all the arthropods originally applied to the vertebrate animal are not9 recovered, the arthropod counts could be subject to measurement error in counting, which more likely would be an issue of undercounting. Bias often becomes apparent when more than 5%-10% of the data are missing. If the extent of undercounting varies considerably between the control and treated groups, or if the application of the treatment leads to very small counts in the treated group (e.g., 0 to 5 pests remain on the animal after application of treatment), the efficacy assessment could be biased. The Panel recommended that measurement error in counting should be investigated by the Agency or taken into account in the modeling framework as discussed in more detail in the response to Charge Question #9.  PG 66
The Agency did not include measurement of error in counting in the models because we do not have an estimate for measurement of error in counting. However, counting live pests is considered the most accurate way to indirectly account for pests.

Live pest counts on the animals are used for general mortality claims for topical/collar products as well as for shampoo products.

Utilizing live counts should have no different bias or effect on Abbott's formula being used in the shampoo tests than what is in the basic flea and tick sections.
308
a. Given the discussion above, assessing live arthropods in the bathing tub is an adequate and practical endpoint for determining mortality. However, the Panel noted that the practicality of this recommendation includes the considerations discussed below.

Tick placement for pesticidal shampoo products should follow the same suggestions previously made for mortality and repellency studies. Ticks should be placed on Day-2 (not Day-1) to permit adequate attachment time before the treatment is applied.

Any live pests found in the plugged tub should be added to the total live arthropod counts. The vertebrate animal should be allowed to fully dry for 24 hours before counting the remaining fleas or ticks. Fleas are, however, more likely to be washed off than ticks, while ticks are more likely than fleas to remain attached to a vertebrate animal and not removed by the act of shampooing. Therefore, the Panel recommended that the EPA guidelines should use different approaches to count the fleas and ticks remaining on the vertebrate animals. A fine-toothed comb should be used to recover fleas; ticks still attached to the vertebrate animal may be discovered by carefully palpating the animal.  PG 66-67
The guideline has been revised to recommend infesting with ticks on day -2 instead of day -1.

The guideline has been revised to allow counting of pests on the animals after the shampooing at up to 24 hours for fleas and 48 hours or ticks.

All live pests found in the plugged tub that are still alive after the alloted time are added to the total live arthropod counts as well.
309
a. The Panel noted that more specific guidelines may be needed for how to assess the mortality of fleas or ticks collected after being washed off or recovered from the vertebrate animal after the shampoo treatment as this can be a difficult endpoint to assess. The determination of tick mortality (whether the tick is attached or unattached to the vertebrate animal) can be evaluated by breathing on the pests and looking for movement, but there may be a delayed response.

Fleas and ticks may not die immediately after they are removed from a vertebrate animal, but these pests might die soon after being removed. The death of the removed arthropods may or may not be due to the pesticidal ingredient in the test product. The death of fleas and ticks after detachment from a vertebrate animal may occur because they no longer have access to blood or are not maintained in adequate environmental conditions. Fleas and ticks need appropriate levels of humidity and temperature to ensure that their mortality is not due to environmental factors alone. Conversely, the pest may be moribund and not recover due to the pesticidal treatment. The mortality of the recovered arthropods should be evaluated after the pest is removed from the vertebrate animal to determine if the pest will recover or die from treatment within a short timeframe. Due to the vulnerability of the arthropod while off the animal, the mortality of fleas should be evaluated within 24 hours. Because ticks might take up to 48 hours to recover, the mortality of ticks should be evaluated within a time window of 24-48 hours.  PG 67
The guideline has been revised to add that after the pest holding period, pests should be considered dead if probed with no movement.

The guideline has been revised to allow counting of pests on the animals after the shampooing or in petri dish after being removed from the water up to 24 hours later for fleas and 48 hours or ticks.

To mitigate flea or tick deaths after removal from the tub due to environmental causes, the following statement has been added to the guideline: "all fleas and/or ticks found in the tub should be removed and moved to a clean container to dry in environmental conditions conducive to their survival for up to 24 hours for fleas and 48 hours for ticks."
310
a. Soaps have a mortality factor against fleas and ticks. Shampoo may remove the waterproofing ability of the pest and make the arthropod more vulnerable to desiccation. Therefore, the Panel recommended that the guidelines should specify that the appropriate control to compare against the pesticidal shampoo is the shampoo without the active pesticidal ingredient.  PG 67
The guideline has been revised to remove the recommendation to use a non-pesticidal (vehicle) control for shampoos. See responses to comment #s 311 & 317 for more information.
311
a. The use of a control shampoo without the active ingredient is needed to determine whether the pesticidal agent had a greater effect than that of the shampoo alone.  PG 67
The guideline has been revised to remove the recommendation to use a non-pesticidal (vehicle) control for shampoos since the product formulation is registered and not the active ingredient. See response to comment #317 for more information.
312
a. Each vertebrate animal should only be shampooed according to the product claims, and not continuously.  PG 67
This question is for shampoo products that are claiming residual pesticidal claims. Therefore, animals are only shampooed according the label directions. Treatment day is on day zero of the study.
313
a. Shampoos do not usually have long-term residual efficacy against fleas and ticks. A study by Beugnet et al. (2012) indicates that weekly shampooing of dogs reduced the number of fleas compared to controls, but the overall reduction in flea number never rose above 79.2%, which would not meet the EPA required efficacy of 90.0%. The Beugnet et al. (2012) study also showed that missing a weekly shampoo resulted in a decrease in efficacy from 68.2% to 34.8%. To assess residual activity of the shampoo on the vertebrate animal, it is important to wet and bathe the vertebrate animal, and then allow the vertebrate animal to fully dry for 24 hours before re-infesting with fleas or ticks on the following day. Counts for fleas should be made on the day following re-infestation and counts for ticks should be made two days after re-infestation to confirm that the active ingredient has not been removed from the animal.  PG 67-68
The guideline currently reflects this comment. If a registrant wants to make residual claims, they should re-infest according to the schedule listed in the above sections. The guideline has been revised to indicate that the animals should be dry before reinfestations.
314
a. The Panel recommended that the guidelines should specify that a vertebrate animal should only be infested when their fur is dry (not with wet fur). In addition, it is not practical to try to comb count a vertebrate animal with wet fur.  PG 68
Both of these suggestions have been incorporated into the guideline.
315
b. The Panel was not aware of any alternative methods that are adequate for determining mortality of fleas and ticks from shampoo products. However, the Panel recommended that the Agency guidelines be open to reviewing and including alternative methods for determining mortality of pests for shampoo products as these new alternative methods proceed through development and become available.  PG 68
The guideline indicates that novel methods may be discussed with the Agency or submitted as a protocol.
316
b. In addition, the Agency should review the process that the U.S. Food and Drug Administration (FDA) uses to assess and approve topical insecticidal products available as prescription and over- the-counter drugs used on people and on animals. The FDA has approved insecticides as topical lotions and shampoos for human use against lice and topical insecticidal solutions for pets. Some useful assistance could be found by reviewing FDA protocols for approval of these drugs.  PG 68
The FDA does not have a published guideline to review on effectiveness of shampoo products and instead follows the EMA guideline, which was cited in this guideline.

Public Recommendations (docket ID # in parentheses): n/a

INSECTICIDAL SHAMPOO PRODUCTS (SECTION (M))

Charge Question 23
Typically, efficacy testing for pet products is conducted with a treated group and an untreated control group. To determine efficacy of shampoo products, the mechanical removal of arthropods during the shampoo process should be considered. Efficacy of a product would therefore be considered in relation to a group treated with non-insecticidal shampoo. In this guideline we propose a non-pesticidal/non-medicated shampoo as the control when testing insecticidal shampoo products.
a. Please discuss the value of using a non-pesticidal/non-medicated shampoo as the control
    i. Does this provide valuable additional information about efficacy beyond an untreated control?
    ii. If so, please provide an appropriate methodology for the comparison of treatment vs. control groups, including references.
b. Should an untreated control also be provided for comparison with the non-pesticidal shampoo? If so, please describe the most appropriate untreated control (e.g., water only, massaging the animal). What is the value of the untreated control and is the informational value of an additional treatment group offset by the increased number of vertebrate animals used for testing?
c. If an untreated control group is included, how should efficacy be determined? Which control group (the non-insecticidal shampoo control or the untreated control) should be compared with the treated group to determine efficacy?

Panel Recommendations:
317
Charge Question #23 focuses on the utility of non-pesticidal/non-medicated shampoo control for determining the efficacy of its pesticidal/medicated shampoo counterpart. Unlike the inert ingredient(s) in other (i.e., non-shampoo) pesticidal products, the ingredients of shampoo are not "truly inert." Specifically, shampoo typically contains a variety of ingredients; these may include water, surfactants (surrounds dirt and oil so water can rinse them away), foaming agents (provide the "suds"), an acid (to balance pH), silicones (create smoothness and add shine), polyquaternium (like a fabric softener), panthenol (fatty alcohols and oils for hydration), fragrances, and preservatives. Since arthropod ectoparasites are relatively small and have large surface area to body weight ratios, anything that disrupts their waterproofing (epicuticular lipids, especially hydrocarbons) could cause desiccation and knockdown even in a relatively high humidity environment. Also, the mechanical action of shampooing can remove the ectoparasites from their hosts.  In other words, the non-pesticidal/non-medicated shampoo may kill the ectoparasites or hamper their ability to adhere onto their hosts. In fact, non-pesticidal/non‐medicated shampoo was shown to reduce the flea count on dogs (Beugnet et al., 2011).  However, in the same study by Beugnet et al. (2011), unlike the pesticidal/medicated shampoo employed, weekly shampooing was needed to maintain the flea count reduction (i.e., continuous control), and the efficacy of shampooing alone on the flea count reduction was shown to be below the performance standard (i.e., >90%) required by different regulatory agencies including the U.S. EPA. Hence, the value of a pesticide in a pesticidal/medicated shampoo product is to provide the continuous control (an important attribute of the product's commercial viability) that cannot be achieved through "treatment" by shampooing only. Accordingly, the utilities of the shampoo control are to control for the short term (i.e., non-continuous) pesticidal action of shampoo "inert" ingredients, as well as to assess the efficacy of pesticidal/medicated shampoo for providing the continuous control. Following a lengthy discussion, the general consensus among most Panel members, except for one Panel member (please see the last paragraph), was that a non-pesticidal/non-medicated shampoo control (i.e., formulation blank) should be used for determining the efficacy of the corresponding pesticidal shampoo product.  PG 69
The use of a non-pesticidal (vehicle) control has been changed to an untreated/negative control for shampoo products, since the Agency typically recommends testing on the end use products.
318
As mentioned previously, a completely untreated control (i.e., host receiving no shampooing treatment) is not an appropriate control for the efficacy determination of pesticide in a pesticidal/medicated shampoo product because it does not control for the short-term pesticidal effect of shampoo "inert" ingredients. Similarly, control with water only or messaging "treatment" (i.e., placebos) provides no more information than the non-pesticidal/non‐medicated shampoo control (Taenzler et al., 2016).  Accordingly, the inclusion of such untreated control(s) will only increase the number of animals used.  PG 69
See response to comment #317 above.
319
Most Panel members concluded that for evaluating the performance of the pesticide in a pesticidal/medicated shampoo product, only two groups are needed: a treatment group with the pesticide added to the pesticidal/medicated shampoo product and a shampoo control group with the formulation blank. Because the data collected are counts, a nonparametric test such as Mann-Whitney rank-sum test can be used to compare the treatment versus the control groups (Glantz, 2012).  In conclusion, the efficacy of pesticidal shampoo product should be determined relative to a non-pesticidal shampoo, or as close a formulation as possible to the candidate pesticidal shampoo.  PG 69
See response to comment #317 above.
320
As mentioned above, one Panel member, suggested that the decision on "what is the appropriate treatment of control group?" depends upon the question(s) to be answered and the product claims; specifically, what is (1) the efficacy of the product as sold (shampoo with the pesticide added), (2) the efficacy of the addition of the pesticide to a non-medicated shampoo base, and (or) (3) the need to know the efficacy of each separately - the non-medicated shampoo base alone and the pesticide alone. To address these questions, this Panel member proposed the following experimental designs (in the same order as the questions raised) for the product efficacy testing: (1) the use of water as the control group and the shampoo containing the pesticide as the treatment group, (2) the use of base shampoo without the pesticide for the control group and the shampoo containing the pesticide as the treatment group, and (3) the use of a three group design: water, the base shampoo and the base shampoo containing the pesticide. The last may need additional animals since two effects are being tested.  PG 69-70
A three-part design would increase the number of animals required for the test, and the Agency believes the revised study design, using an untreated control, is sufficient to determine shampoo product efficacy.

Public Recommendations (docket ID # in parentheses):
321
"Application methods. Insecticidal shampoo evaluations should be compared against a non-treated group or a non-treated group and a non-insecticidal shampoo group. This is because noninsecticidal shampoo may have an effect on some of these arthropods. Any effect on the insect population will translate into false positives in the data evaluation. If it must be considered to have a "placebo" than it is suggested that a water only control supplant the placebo. The trial must still include a totally untreated control group to evaluate for confounding of results. This results in a requirement for additional test animals... Change in shampoo testing by use of inert control, will cause issue since a non-medicated shampoo will also wash off some pests making the statistic much more difficult to show statistical significance. It is also a change to an unreal life situation" (0008)
The guideline has been revised to compare the treated shampoo against an untreated control only.
322
"Testing using a non-pesticidal shampoo as a control results in a test for the effect of the pesticide only, not the pesticidal shampoo. The product that the consumer buys is the pesticidal shampoo, so the pesticidal shampoo should be tested against an untreated control" (0033)
The guideline has been revised to incorporate this comment.
337  -  out of order
"Representative sampling. Including a non-medicated, non-insecticidal shampoo in protocols for insecticidal shampoo testing will artificially impact pest counts and efficacy evaluation. Under field conditions, the user would not shampoo an animal with a non-insecticidal shampoo and then an insecticidal shampoo." (0008)
The guideline has been revised to incorporate this comment.

SIMULATED ENVIRONMENTAL CONDITIONS (SUNSHINE, WATER EXPOSURE AND SHAMPOOING CLAIM) (SECTION (N))

Charge Question 24
Discuss whether the timing for infesting vertebrate test animals after water exposure are adequate.
a. Are the number of infestations presented in the guideline adequate to support bathing and water immersion claims?
b. Discuss appropriate timing of infestations after vertebrate test animals are immersed in water.
c. Are the methods for determining exposure to sunlight adequate? If not, please provide detailed methods for evaluating the effects of exposure to sunlight along with supporting references.

Panel Recommendations:
323
The Panel recommended that the reinfestation be specified as 24 hours after each exposure to water or bathing (e.g., bathing at day 6 with reinfestation at day 7, bathing at day 13 with reinfestation at day 14, etc.). In this way the animal will be dry prior to the reinfestation, even for animals with thick coats of fur.  PG 70
The guideline already states that "when infestations follow shampooing/wetting they should occur within 48 hours of shampooing/wetting. Animals should be dry before pest infestations." This should provide the lab some flexibility, while explaining the expectation.
324
The Panel recommended the use of a "non-medicated" shampoo as the control, because this represents a baseline of control that a pet owner could achieve on their own without a shampoo containing a pesticide. The Panel did not recommend the use of the additional water exposure control, as the value of such studies does not justify the number of vertebrate animals needed for this.  PG 70
These tests are intended to demonstrate that a product used to treat an animal will still work after the animal has been bathed or exposed to water. The untreated group that has been bathed or exposed to water is the untreated control and is handled in the same manner as the treated group. Therefore, those test animals are still needed to serve as controls. No changes will be made to the guideline.
325
a. Yes, the Panel found that the number of infestations presented in the guideline are adequate to support bathing and water immersion claims.  PG 70
No changes to the guideline are needed.
326
b. The Panel recommended that the reinfestation be specified as 24 hours after treatment. In this way the vertebrate animal will be dry prior to the reinfestation, even for animals with thick coats of fur.  PG 70
See response to comment #323.
327
c. It is difficult to state unequivocally if the methods for determining exposure to sunlight are adequate or not. The current protocol does not really clarify if the product will be stable in sunlight. The vertebrate animals will be outside for part of the day but will be provided shade. Will the animals go into the sunlight? How long will they go into the sunlight? The UV index differs geographically, and this would potentially impact the results. Monitoring the sunlight each animal receives (using an on-collar monitor) would probably be necessary to standardize exposure and back up claims that a product continues to work after some specified amount of sunlight exposure. The Panel recommended that if the current approach is used, that the host be exposed to sunlight for two hours per day. This would potentially offer some standardization, although variation in weather may make such a regulation difficult or impossible to achieve. It would be of some additional value if the daily weather (average daily temperature, high temperature, low temperature, sunshine, rain, etc. was also recorded). While such information may not alter the approval process, these variables would help understand differences that might occur between studies.  PG 70-71
The suggestions for animals to be preferably exposed to direct sunlight for an average of 2 hours a day has been added as well as requesting daily records of the amounts of sunshine and rain to be submitted.
328
The Panel noted that under the current proposed guidelines, vertebrate animals will spend prolonged periods outdoors and that some of them (i.e., the controls) could pick up additional or new pests while outdoors. As such, it would be valuable if the total number of pests were reported on the controls and treated animals at the end of the study, not just the number of pests that the animal was infested with.  If the outdoor activity of the pets results in accumulation of additional pests of the same species that they were infested with, the number of control animals might need to be increased to get a better assessment of this.  PG 71
 The "infestation period" of these studies are still conducted in a lab, therefore the animals should still be kept separated in outdoor runs during that time as indicated in the mortality testing. However, all animals in the study will have counts conducted, therefore, if control animals picked up" new infestations" it would be reported, and the study would be questionable or invalid. No changes to the guideline are needed.
329
The Panel also considered using artificial sunlight on caged animals. This has potential for standardization of the "sunlight" exposure but requires protection of the animal's eyes and raises other animal welfare concerns. As such, the Panel did not recommend this approach.  PG 71
No changes to the guideline are needed.
330
The Panel also suggested that EPA consider claims for sunlight stability using a post-approval field study with client-owned dogs under real world conditions with natural pest infestations in different geographic areas of the US and with a significant portion of the enrolled dogs in the study that spend at least half of their time outdoors with exposure to sunlight (collar monitors to record sunlight would be beneficial).  PG 71
While the use of post-approval field studies with client-owned dogs would decrease the need of lab animals for this specific test, previously submitted field studies have not yielded useful data. Issues included inadequate or inconsistent pest pressure, lack of appropriate controls, and inaccurate pest counting by owners. However, protocols for field studies may be submitted if desired.
331
Given the difficulty in understanding the amount of sunlight the animal is exposed to, the Panel recommended consideration of non-vertebrate testing of treated fur.  The treated fur could be exposed to sunlight and the residue of the active ingredient determined after different days or weeks of exposure to sunlight.  PG 71
This is a recommendation to consider in the future, however it is currently unknown how a product breaks down by sunlight in the fur vs. the lipid layer of the skin, where the products are often spread and regenerated into the fur. No changes to the guideline will be made.
332
Suggested editorial changes:
The Panel noted that in section n(1)(i) for products with efficacy claims of 2-4 weeks, animals should be bathed on Days 6 and 20 so infestations can occur on appropriate Days 7 and 21.  The rationale is that the dogs that are wetted should be allowed to dry for ca. 24 hours prior to being re-infested the next day.  PG 71
The guideline has been revised based on the suggestion.
333
Suggested editorial changes:
Section (n)(1)ii. The Panel suggested the following revision "fur and skin through submerging (except the head) or showering the animal"  PG 71
The guideline has been revised based on the suggestion.

Public Recommendations (docket ID # in parentheses):
334
"General testing for bathing & water exposure claims. Separate considerations should be given to collar formulations to be evaluated at 7 days after the shampooing/wetting. These collars must be allowed enough time to re-release an efficacious amount of active ingredient and for that active ingredient to redistribute across the surface of the animal's body. The current proposed 48-hour challenge does not allow for this. This is significantly different from what has been recently accepted for shampooing/wetting claims for collars. Prior EPA approvals of shampooing/wetting claims for collars have allowed a seven-day interval between washing and re-infestation, this is a reasonable amount of time for the redistribution of active ingredients." (0008)
The purpose of the study is to demonstrate that the product still works after bathing or water exposure. Allowing 7 days for re-evaluation would be similar to reapplying the product and waiting for it to begin working again. If the product does not work within 48 hours of wetting or shampooing, it should not be considered "water-proof" or "shampoo-proof."
335
In section n(1)(i) for products with efficacy claims of 2-4 weeks animals should be bathed on days 6 & 20 so infestations can occur on appropriate days 7 & 21" (0012)
The guideline has been revised based on the suggestion.

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