Housing and dustbathing effects on northern fowl mites
(Ornithonyssus sylviarum) and chicken body lice
(Menacanthus stramineus) on hens
C. D. MAR T I N and B. A. MUL L E NS
Department of Entomology, University of California Riverside, Riverside, CA, U.S.A.
Abstract. Hen housing (cage or cage-free) did not impact overall abundances
of northern fowl mites, Ornithonyssus sylviarum (Canestrini & Fanzago)
(Acari: Macronyssidae), or chicken body lice, Menacanthus stramineus (Nitzsch)
(Phthiraptera: Menoponidae). Cage-free hens received a dustbox with sand plus
diatomaceous earth (DE), kaolin clay or sulphur. Weekly use varied from none to
100% of hens; 73% of hens used the dustbox at least once. Ectoparasite populations
on dustbathing hens (users) were compared with those on non-user cage-free and caged
hens. All materials reduced ectoparasites on user hens by 80–100% after 1 week of
dustbox use. Diatomaceous earth and kaolin failed to reduce ectoparasites on non-user
hens, and ectoparasites on user hens recovered after dustbox removal. A sulphur dustbox
eliminated mites from all hens (including non-users) within 2–4 weeks. Residual
sulphur controlled mites until the end of the experiment (up to 9 weeks), even after
the dustbox was removed. Louse populations on hens using the sulphur dustbox were
reduced in 1–2 weeks. Residual sulphur effects were less evident in lice, but the use
of a sulphur dustbox by a higher proportion of hens extended louse control to all
hens. This is the first experimental study to show that bird dustbathing in naturally
and widely available dust materials (particularly kaolin) can suppress ectoparasites
and thus the behaviour is probably adaptive.
The northern fowl mite (NFM), Ornithonyssus sylviarum,
and the chicken body louse (CBL), Menacanthus stramineus,
are primary ectoparasites of poultry (DeVaney, 1978; Axtell
& Arends, 1990). Current control tactics consist primarily
of direct pesticide treatments to the host feathers and, in
some cases, treatments of the hen housing environment (i.e.
litter, nestboxes) to kill any ectoparasites that linger between
flocks. Unfortunately for producers, few synthetic chemicals
are available for mite control and resistance is problematic
(Mullens et al., 2004a). Further, although chemical control
and population dynamics of NFM and CBL have been
studied extensively in cage settings (Matthysse et al., 1974;
Arthur & Axtell, 1983; Mullens et al., 2009), scientific studies
examining mite and louse population dynamics and control in
hens held in cage-free conditions are lacking.
Animal welfare issues also are becoming increasingly important
in poultry systems (Rodenburg et al., 2008; Lay et al.,
2010). This has resulted in mandated changes in poultry
housing, such as the Welfare of Farmed Animals Regulations
Act of 2000 (http://www.opsi.gov.uk), and California’s
Proposition 2 (Standards for Confining Farm Animals)
(Sumner et al., 2008). In general, changes require moving
away from typical wire (‘battery’) cages, and providing hens
with greater space and/or altered or more complex cage
designs. Such changes could result in a potentially serious
economic disadvantage in states such as California relative
to other states lacking those requirements (Sumner et al.,
Pest control in the poultry industry must be tailored to the
different production systems (e.g. cages, enriched cages, cagefree,
free range). Notably, cage-free flocks cannot be treated
as easily as hens in cages, which can be treated using highpressure
chemical spraying from below. Even this method,
however, can result in incomplete coverage (Axtell, 1999).
Dustbathing behaviours have been described in chickens
(van Liere, 1991), wrens (Hendricks & Hendricks, 1995) and
quail (Borchelt et al., 1973), but few studies have experimentally
addressed the adaptive value of the behaviour. Literature
reviews have discussed the proven significance of dustbathing
for feather maintenance and have hypothesized that it reduces
ectoparasite numbers, but dustbathing effects on ectoparasites
have never actually been demonstrated experimentally in wild
or domestic birds (van Liere, 1992; Olsson & Keeling, 2005;
Clayton et al., 2010). Earlier studies tested pesticides or biological
agents in dustboxes for ectoparasite control, but individual
hen dustbox use and effects of untreated dust were not well
evaluated (Rodriguez & Riehl, 1957, 1960; Hoffman & Hogan,
1967; Hoffman & Gingrich, 1968).
The present study compared the population dynamics and
abundances of NFM and CBL in cage vs. cage-free settings. It
also explored dustbox use and the self-application of inorganic
dust materials as alternatives to the use of traditional synthetic
pesticides. Finally, the dustbox studies served as the first
experimental evaluation of the potential adaptive value of
dustbathing for ectoparasite suppression using available natural
substrates.
Materials and methods
Experiment design
Studies were conducted at the University of California (UC)
Agricultural Experiment Station next to the UC Riverside
campus, and procedures were approved through the UC
Riverside Institutional Animal Care and Use Committee.
Three trials were conducted between late autumn and late
spring (Trial 1: March–June 2009; Trial 2: October 2009 to
February 2010; Trial 3: March–June 2010). Beak-trimmed Isa-
Brown (Trials 1 and 3) or Hyline Brown (Trial 2) pullets were
obtained from a commercial operator. These breeds are popular
in cage and cage-free systems.
The research poultry houses (Fig. 1) measured 3.8 × 5.8 m,
were screened to exclude the entry of wild birds and
rodents, and equipped with roof sprinklers to maintain internal
temperatures below 33–35 ◦C. Hobo temperature recorders
(Onset Computer Corp., Bourne, MA, U.S.A.) recorded air
temperatures in each house. All hens had ad libitum access
to commercial lay mash in feed troughs and water was
continuously available. Lights inside the houses provided an
LD 16 : 8 h light cycle year-round, as is usual in commercial
production.
Twelve caged hens were held in each of two houses in
suspended wire cages. A rate of two hens per 31 × 31-cm
cage met U.S. United Egg Producer Standards (2010). Cagefree
hens were maintained in each of two houses, divided by
a plastic tarpaulin into north and south halves (3.8 × 2.9 m).
This allowed a dustbox treatment in one half of the house
to be compared with a no-dustbox treatment among cage-free
controls in the other half of the house (12 hens per treatment).
Cage-free houses contained eight nestboxes and perch space.
Each nestbox was about 0.1 m2 in area and was elevated about
0.8 m above the ground. Straw (2–4 cm deep) covered the
concrete floor At 19 weeks, 18 weeks and 20 weeks of age (in Trials 1–3,
respectively), each hen was infested with either 20–30 mites
(mostly adult females) or 20 lice (mostly adults, mixed sexes).
Mites were collected from a colony originally established in
2003. Mites were aspirated into capillary pipettes from source
hens and transferred to the vent area feathers (in an 8–9-cm
diameter zone extending from the base of the legs to the posterior
end of the cloaca) and skin of test hens, using a fine paint
brush. Lice (from a colony originally established in 2005) were
collected from source hens by inserting the hose from a carbon
dioxide (CO2) tank into the vent and abdominal feathers. This
lightly anaesthetized the lice, which were then shaken from the
abdominal feathers into a small bucket from which they were
transferred using a fine brush to the abdominal feathers of their
new hosts. One caged hen group and one cage-free hen group
were infested with NFM. Likewise, one caged hen group and
one cage-free hen group were infested with CBL.
All hens were identified as individuals by leg-banding one
of the two hens per cage and by using unique combinations
of coloured bands on the two legs of cage-free birds.
Ectoparasite counts were conducted weekly for 14 weeks
for NFM and 15 weeks for CBL. Inspections for NFM and
CBL were performed on different days to prevent crosscontamination.
Only the area near the vent was inspected
visually for NFM, using a scoring system (Arthur & Axtell,
1982) modified by Owen et al. (2009). Briefly, a score
of 0 = no mites seen, 1 = 1–10 mites, 2 = 11–50 mites,
3 = 51–100 mites, 4 = 101–500 mites, 5 = 501–1000 mites,
6 = 1001–10 000 mites, and 7 = >10 000 mites. A plus (+)
or minus (−) (except for a score of 7) was used to indicate a
population level in the upper or lower 20% of the respective
range.
In inspections for CBL, the chicken vent region was
examined as described for NFM. The area under each wing,
measuring approximately 8–10 cm in length and 5–7 cm in
width, consisting of the axilla and three feather partings down
the side of the body (separated by about 1.5–2.0 cm) was
also observed. Lice from the vent and under-wing areas were
summed.
Dustbox studies
Dustboxes (Fig. 2) had a 1.9-cm-thick plywood bottom, 2.5-
cm-thick wood sides and measured 65 × 65 × 20 cm. A 5-cmwide
wooden lip helped keep the particle mixtures in the box.
Washed playground sand (Premium Play Sand
; Quikrete
Companies, Inc., Atlanta, GA, U.S.A.) consisted of silica
particles of 0.5–1.5 mm in diameter. A 9 : 1 ratio by weight
of sand to dust [by volume: about 50 : 50 for diatomaceous
earth (DE) : sand and kaolin : sand, and about 25 : 75 for
sulphur : sand] was used in all three trials. The starting mixture
consisted of 10 800 g of sand and 1200 g of 100% DE
(Organic D/Earth; Thomas Laboratories, Inc., Tolleson, AZ,
U.S.A.), 95% kaolin (Surround WP; Engelhard Corp., Iselin,
NJ, U.S.A.) or 90% wettable sulphur (Yellow Jacket; Georgia
Gulf Sulfur Corp., Valdosta, GA, U.S.A.), which filled the
dustbox to a depth of 3.5–5.0 cm. A small amount (0.16% by
weight) of fluorescent dust (FD; ‘arc yellow’ or ‘signal green’;
Day-Glo Color Corp., Cleveland, OH, U.S.A.) was added as
a dust marker (Hall et al., 1981) to estimate dustbox use by
individual hens. By shining a portable longwave ultraviolet
flashlight (WARD’S Natural Science, Rochester, NY, U.S.A.)
into the abdominal feathers under subdued ambient light, we
were able to assign a dustbathing score based on a combination
of the fluorescence level and overall visual ‘dustiness’ of
the hen. Scores were: 0 = no dust; 1 = trace dust (<25%
coverage of vent area, usually <10 individual FD particles);
2 = moderate dust (FD covering 25–50% of inspected vent
area), and 3 = high dust (FD covering 50–100% of inspected
vent area and particles often made airborne when parting
through feathers).
Each trial ran for 15 weeks; caged (no-dustbox) hens
were checked for ectoparasite numbers weekly. Cage-free
mite hens received a dustbox for weeks 4–8 [north end
(N)] and weeks 8–12 [south end (S)]. Cage-free louse hens
received a dustbox for weeks 5–9 (N) and 9–13 (S). This
provided experimental (dustbox treatment) replication with
two groups of hens for each ectoparasite. Other than those
4-week periods, the cage-free hens did not have access to
a dustbox. Thus, mites and lice on cage-free (N) hens were
given 4 weeks and 5 weeks, respectively, to reproduce before
hens were given access to a dustbox. Mites and lice on cagefree
(S) hens were given 8 weeks and 9 weeks, respectively,
to reproduce before hens were given a dustbox. Each cagefree
hen group was given a single dustbox for a 4-week
period, after which the box was removed. Dustboxes in
Trial 1 (DE) were replenished every other week with 50% of the starting material amounts; dustboxes in Trial 2 (kaolin)
were replenished weekly with 25% of the starting material
amounts. This replenishment was carried out to keep the
dustboxes effective because they appeared to lose the fine dust
material as the chickens scratched and kicked it out during
dustbathing. Trial 3 dustboxes were never replenished because
the effectiveness of the sulphur quickly became evident and
replenishment seemed unnecessary.
Data analysis
Statistical analyses were carried out using Minitab Version
14 (Minitab, Inc., State College, PA, U.S.A.). For parametric
analyses, the normality of each dataset was assessed using
the Ryan–Joiner test. One-way analyses of variance (anovas)
were used within a trial to generate weekly means for graphing and to compare weekly ectoparasite (NFM or CBL)
populations between hen groups. Housing effects (pre-dustbox)
data were also subjected to repeated-measures anova.
Regression slopes (abundance vs. time) were utilized as a
general indicator of population trends after dustbox removal
(mainly for DE and kaolin treatments). Slopes for the ‘during’
dustbox phase were based on a 5-week period, including the
ectoparasite level at the beginning of dustbox placement, and
those for each week until dustbox removal. Slopes used for the
‘after’ dustbox phase were based on periods of 3 weeks (cagefree,
S) to 5 weeks (cage-free, N), including the ectoparasite
level at the time of dustbox removal. This spanned the period
of 2–4 weeks post-dustbox removal. The effects of dustbox
use on ectoparasites were analysed by contrasting populations
on ‘users’ (FD scores of 2 or 3) with those on ‘non-users’
(FD scores of 0 or 1), employing two-sample t -tests. To
compare relative, short-term ectoparasiticidal effects of the
three materials, converted mite scores or louse numbers on
dustbathing hens (from week 0 to week 1 of dustbox use) were
compared with those on caged controls. Using the Henderson
& Tilton (1955) formula to calculate corrected percentage
control, the relative effects could then be determined by:corrected %
=
⎛
⎝1 −
n in Co before treatment
× n in T after treatment
n in Co after treatment
× n in T before treatment
⎞
⎠ × 100
where n = pest population, T = treated hens, and Co = control
hens.
In these analyses, mite indices were converted to approximate
‘actual’ values (0 = 0, 1− = 2, 1 = 5, 1+ = 8, 2− =
15, 2 = 30, 2+ = 45, 3− = 55, 3 = 75, 3+ = 95, 4− = 150,
4 = 300, 4+ = 450, 5− = 550, 5 = 750, 5+ = 950, 6− =
2000, 6 = 5000, 6+ = 8500, 7 = 20 000) to resolve dustbox
effects.
Results
Housing effects: northern fowl mites
Average weekly mite indices are illustrated in Fig. 3. To
isolate housing effects, these analyses reflect ectoparasite levels
prior to dustbox placement. Scores for NFM at equivalent
stages post-inoculation varied significantly between trials
(F = 13.41; d.f. = 2, 470; P <0.001). Therefore, trials were
examined individually.
Northern fowl mite population growth did not differ overall
between cage and cage-free treatments (weeks 1–
in any of
the three trials (F ≤ 2.58; d.f. = 1, 145 or 1, 154; P ≥ 0.123).
Average weekly scores in the two housing types differed
occasionally (Fig. 3), but the variations were relatively minor
and not consistent. Differences between the cage-free (N)
treatment vs. the cage treatment were seen in Trials 1 and 2
(F <8.91; d.f. = 1, 66; P <0.03), but not in Trial 3 (F =
0.01; d.f. = 1, 66; P = 0.913). When weekly differences were
seen between the north and south cage-free flocks, the occurred in the hotter weather conditions of Trial 1. Degree
days (DD) above 25 ◦C were highest in Trial 1 (DD = 63.
and lower in Trials 2 and 3 (DD = 12.3 and DD = 7.3,
respectively). Both housing types allowed large numbers of
mites to develop on hens.
Housing effects: chicken body lice
Average louse numbers per hen prior to dustbox placement
differed significantly among the three trials (F = 25.32;
d.f. = 2, 540; P <0.001) (Fig. 4) and therefore trials were
examined separately. Louse abundance in cage vs. cagefree
(S) treatments did not differ significantly (weeks 1–9)
in any of the three trials (F ≤ 2.75; d.f. = 1, 172 or 1, 176;
P ≥ 0.112). Similarly, overall cage and cage-free (N) numbers
(weeks 1–5) did not differ significantly in any of the three
trials (F ≤ 1.44; d.f. = 1, 88; P ≥ 0.242). The rare weekly
differences were inconsistent. Both housing types allowed large
numbers of ectoparasites to develop on hens.
Dustbox studies: general trends
Figures 5 and 6 illustrate average on-host population trends
for NFM and CBL, respectively, in Trials 1–3 and depic three phases in cage-free treatments: (a) increasing ectoparasite
loads to high levels prior to dustbox placement; (b) ectoparasite
decline caused by hen dustbathing, and (c) general recovery
or stability of mite and louse populations after the removal
of DE and kaolin dustboxes. In general, standard errors of
mean ectoparasite numbers were higher when dustboxes were
present, reflecting differential dustbox use among hens.
Overall, mites and lice were reduced most effectively on
dustbathing hens, whereas non-user hens maintained moderate
to high levels of infestation (except for non-user hens in
houses with a sulphur dustbox). Figures 7 and 8 show average
ectoparasite levels on dustbox users compared with non-users
from week to week, and also indicate how many hens were
dustbox users in that week.
Over time, the straw on the cage-free floors degraded into
finer particles as the hens dustbathed, walked and scratched
in it, and straw particles became mixed with manure and
feed. As well as dustbathing in the boxes, hens would do
this in the litter before, during and after dustbox placement.
Based on the sieving of four random 1-liter samples at the
end of Trial 3, the particle size distribution [mean ± standard
deviation (SD)] was as follows: particles measuring >2.4 mm
accounted for 63 ± 16% of samples; particles of 0.6–2.4 mm
for 31 ± 10% of samples; particles of 0.25–0.6 mm for 5 ± 5%
of samples, and particles of <0.25 mm for 1 ± 1% of samples.
Notably, dustbathing in the litter apparently had no impact
on ectoparasite populations on cage-free hens; their numbers were similar to those on caged hens that were denied any
dustbathing opportunities. There were also obvious effects of
experimental dustbox use on ectoparasites when the boxes were
available (see below). These effects across time were clear
when contrasted with effects during periods when cage-free
hens could dustbathe only in litter.
Dustbox studies: diatomaceous earth
After 1 week of fresh dustbox access (week 4–5), average
mite counts (converted from NFM indices) on the six cagefree
(N) dustbathing (user) hens declined by 86% (75–100%
per hen) (Fig. 7A), whereas average mite counts on the six
cage-free (N) non-dustbathing hens increased by 30%. Mites
on cage-free, untreated hens (S) dropped by 61% and mites on
caged hens increased by 4% (Fig. 5A). User hen mite indices
were significantly lower than indices for non-user hens for
weeks 5, 6 and 8 (P ≤ 0.05) (Fig. 7A).
In a replication allowing fresh dustbox access with the
second hen group (week 8–9), mite numbers on 10 cagefree
(S) dustbathing hens declined by 79% (Fig. 7B), whereas
numbers on cage-free (S) non-user and caged hens dropped by
33% and 55%, respectively (Figs 7A and 5B, respectively).
Hens using the dustbox had significantly fewer mites than
non-user hens at weeks 11 and 12 (P ≤ 0.01) (Fig. 7B). Mite score regression slopes post-dustbox removal in Trial 1
did not differ significantly from zero (P ≥ 0.114). Mites on
caged hens during these periods also exhibited slight overall
negative regression slopes, but, again, these did not differ
significantly from zero (P ≥ 0.139).
After 1 week of dustbox access (week 5–6), average cagefree
(N) louse counts declined by 88% on the three dustbox
user hens and increased by 10% on non-users (Fig. 8A). In
the same period, louse populations on caged and untreated
cage-free (S) hens increased by an average of 50% and 40%,
respectively (Fig. 6A, C). Average louse counts on user hens
were significantly lower than on non-user hens (P ≤ 0.05)
(Fig. 8A) in every week of dustbox access.
In a replication with the second hen group (week 9–10),
average louse counts on nine of 11 cage-free (S) dustbathing
hens declined by 96%, whereas average counts on non-users
declined by 26% (Fig. 8B), and counts on caged controls
declined by 14% (Fig. 6A). No significant differences in louse
numbers were detectable between user hens and the very few
non-user hens in weeks 10–13 (Fig. 8B).
Regressions of louse numbers over time in the weeks
following DE dustbox removal produced positive slopes in
both treatments, although neither differed significantly from
zero (P ≥ 0.251) (Fig. 6B, C). Similarly, lice on caged hens exhibited no significant slope deviation during this recovery
period.
Dustbox studies: kaolin
After 1 week of fresh dustbox access (week 4–5), average
mite abundance on five of 12 cage-free (N) dustbathing hens
declined by 95%. Mite numbers on seven of 12 cage-free (N)
dustbox non-user hens increased by 10% (Fig. 7C). Mites on
untreated caged and cage-free (S) control groups increased by
230% and 370%, respectively (Fig. 5A, C).
In a replication with the second hen group (week 8–9), mite
numbers on six of 12 cage-free (S) dustbathing hens declined
by 90% after treatment and those on non-users remained stable
(Fig. 7D). Numbers on caged controls and cage-free (S) nonusers
increased by 80% and 50%, respectively (Fig. 6A, C).
Numbers of mites recovered upon dustbox removal in
Trial 2, as indicated by significantly positive regression slopes.
cage-free (N) scores exhibited a slope significantly greater than
zero (P = 0.003) and increased until mite populations reached
levels approximate to those on caged birds.
Average louse numbers declined by 88% after 1 week of
dustbox access (week 5–6) on six of 12 cage-free (N) dustbox
users but increased by 90% on non-user hens (Fig. 8C).
Meanwhile, louse abundance on caged and cage-free (S)
control groups increased by 70% and 110%, respectively
(Fig. 6A, C).
In a replication with the second hen group, the cage-free (S)
hens in this trial did not accept the dustbox in the first week
(week 9–10) for unknown reasons. By the second week, two
of 12 hens used the resource. Louse numbers on these two
birds were reduced by 73% and 98% (Fig. 8D).
Louse population regression slopes were positive for the
period post-kaolin dustbox removal in both cage-free treatments.
The slope for cage-free (N) louse trends in Trial 2
(post-dustbox) was significantly positive (P = 0.005).
Dustbox studies: sulphur
After dustbox access (week 4–5), mite numbers on three
of the 12 cage-free (N) dustbathing hens declined by 99.9%
(Fig. 7E). Interestingly, mite numbers on nine of 12 cagefree
(N) non-dustbathing hens also declined by 99%. Mite
numbers on caged and cage-free (S) control groups dropped
by 12% and 35%, respectively (Fig. 5A, C).
In a replication with the second hen group (week 8–9),
mite numbers on three cage-free (S) dustbathing hens declined
by 99.9% (Fig. 7F), whereas numbers on caged and cagefree
(S) non-user hens dropped by 19% and 29%, respectively
(Fig. 5A, C). Dustbathing in sulphur was 100% effective in
reducing mites to low levels (scores of <2) after 1 week of
access, and all visible mites were eliminated in a 2–4-week
period (Fig. 5B, C).
Mites did not recover from sulphur dustbox treatments
up to 6 weeks after dustbox removal (Fig. 5B, C), although
relatively few (≤50%) of the hens in each group used the
dustbox on a weekly basis.
After 1 week of dustbox access (week 5–6), lice on the
three cage-free (N) user hens dropped by 100% (Fig. 8E).
Interestingly, louse numbers dropped by 88% on non-user
hens, whereas louse numbers on caged controls increased by
40% (Fig. 6A).
In a replication with the second hen group, lice on
seven cage-free (S) user hens (week 9–10) dropped by 98% (Fig. 8F). Louse numbers on non-user hens and
caged controls dropped by 90% and 47%, respectively
(Fig. 6A).
Louse numbers recovered quickly following sulphur dustbox
treatment when fewer hens used the dustbox (Fig. 6B), but
no recovery was observed up to 2 weeks following dustbox
removal after heavier hen use (Fig. 6C). By comparing ‘before’ and ‘after’ ectoparasite levels on
cage-free hens (user hens only) with ‘before’ and ‘after’
levels on caged hen controls (week 0–1 of dustbox use), the
Henderson & Tilton (1955) formula could be used to calculate
corrected percentage short-term control for DE, kaolin and
sulphur treatments (Table 1). Sulphur was the most active
material. Kaolin effects tended to be more active in the short
term than those of DE, particularly for mites. However, DE was
replenished every 2 weeks, whereas kaolin was replenished
weekly.
Discussion
Housing
Ectoparasite abundances on poultry were not substantially
or consistently affected by housing (cage vs. cage-free). Hens
in both cage and cage-free settings were equally capable of
supporting high ectoparasite loads when untreated with dustboxes
(Figs 3 and 4). Using only caged hens, Hall et al.
(1978), Arthur & Axtell (1983) and Mullens et al. (2000)
observed fewer mites under more crowded cage conditions.
In the present study, mite populations on cage-free hens occasionally
tended to be higher early in infestation than those on
hens in cages, but large or consistent ‘density’ effects on mites
were not seen. Mullens et al. (2010) showed that hens held
in pairs per cage supported about twice as many lice as hens
held singly in cages, presumably because crowded hens were
less efficient in grooming. In the present study, no significant
housing effects on CBL abundance emerged in any trial.
Dustboxes
Hen dustbathing in DE and kaolin dustboxes had very
clear, immediate, negative impacts on mite and louse numbers.
These effects were temporary, however, and depended on
the individual hen’s use of the dustbox. Dustboxes had beensuggested for managing mite and louse infestations on poultry farms at least by the early 1900s (Banks, 1907; Pierce &
Webster, 1909). The few existing dustbox studies evaluated
the effects of synthetic insecticides, rather than documenting
untreated substrate effects on ectoparasites (Rodriguez &
Riehl, 1957, 1960; Hoffman & Hogan, 1967). Hoffman &
Hogan (1967) attributed a lack of louse control to nonacceptance
or monopolization of an insecticidal dustbox, but
did not document dustbox use. Hoffman & Gingrich (1968)
conducted a similar study testing hand vs. dustbox applications
of Bacillus thuringiensis Berliner. They demonstrated little to
no control of lice in dustbox treatments compared with direct
hand applications, but again did not document dustbox use.
Diatomaceous earth and kaolin
Dry formulations of DE and kaolin are relatively nontoxic
to humans, although they do pose some inhalation risks
(Lynch & McIver, 1954; Merget et al., 2002). The presumed
mode of action of these materials is arthropod death by
desiccation, which is achieved either by abrading the cuticle
or absorbing lipids from the cuticle that are important for
managing water loss (Wigglesworth, 1945; Ebeling, 1971).
Ebeling (1973) described kaolin clay as a slightly abrasive
material and DE as both abrasive and sorptive. Cook et al.
(2008) found no actual evidence of abrasion of flour mites
[Acarus siro (Sarcoptiformes: Acaridae)] after exposure to
three DE products for 3–72 h. However, significant amounts
of tridecane, a cuticular lipid thought to be an important
waterproofing agent, were detected on all of the DE products
rinsed off the mites. This suggests the main action of the
dusts, at least on mites, may be through absorption, rather than
abrasion.
Inert dusts, such as DE, are used to treat poultry houses for
Dermanyssus (Acari: Dermanyssidae) in Europe (Mul et al.,
2009; Harrington et al., 2011). Dawson (2004) suppressed fleas
and Protocalliphora (Diptera: Calliphoridae) in tree swallow
nests using DE. Kilpinen & Steenberg (2009) tested DE and
kaolin for control of D. gallinae (De Geer) and found that both
killed mites, but the efficacy of each material decreased with
increasing (75–85%) relative humidity. Based on 24-h miteweight loss studies, kaolin clay resulted in 11.9% loss and DE
exposure in 18.2% loss, compared with 9.5% weight loss in
a control group (Kilpinen & Steenberg, 2009). Despite blood
engorgement by this mite, the dusts may cause water loss faster
than it can be replenished.
Sulphur
Sulphur is a natural component of the environment; it is
generally considered to be safe and is currently the active
ingredient in nearly 300 pesticides (http://www.epa.gov). In
small concentrations (mixed with petroleum jelly), it is used
as a topical treatment for control of scabies mites on humans
(Pruksachatkunakorn et al., 2002). As a dust, however, it is
considered slightly toxic and slightly irritating in terms of
inhalation and eye exposure, and should be kept away from
sparks and flames because it has some ignition potential.
Sulphur has probably been used for mite and louse control
in poultry longer than any other material. Sulphur kills by
direct contact and is thought to have fumigant action at
temperatures >21 ◦C (Ware, 1993), which may be especially
effective within the feather layer. Banks (1907) recognized that
sulphur and lime mixtures could be scattered in a poultry house
for control of mites (D. gallinae) and lice and commented on
the fumigant power of sulphur against lice. Creighton et al.
(1943) tested sulphur for control of poultry lice on White
Leghorns as a feed supplement, a direct dust application and
a soil treatment. They reported that consumption by chickens
was ineffective for control, but that direct and soil treatments achieved excellent control within 1–2 weeks, and that residual
effects were poor when test chickens were later exposed to
heavily infested chickens and were re-infested (Creighton et al.
1943). Furman (1952) showed hand-dusting was effective for
NFM control on hens, and Foulk & Matthysse (1963) achieved
success when dusting caged hens for NFM.
Sulphur was clearly the most potent material tested in our
dustbox studies. User hen groups were most affected, but rather
quickly and over time sulphur had effects on ectoparasites
even on non-user hens. Mites were controlled for ≥9 weeks,
although fewer than 50% of the chickens might use the dustbox
in a given week. In a hen group, lice recovered from sulphur
dustbox treatments when few hens were using the dustbox, but
were severely suppressed for ≤3 weeks on all chickens if more
hens used it. This implies that dustbathing hens were largely
responsible for dispersing sulphur in the environment.
Dustbathing and dustbox use
The most significant factor influencing the value of a dustbox
treatment was the variation in the individual’s use of the box.
About 73% of all hens that had access to a dustbox used
it at some point. Some used it more than others, and some
did not use it at all. In previous dustbox studies, dustboxes
were considered unreliable as a control method for mites and
lice because of differences in dustbox usage among individual
hens (Bishopp & Wood, 1917, 1931; Hoffman & Hogan, 1967;
Hoffman & Gingrich, 1968).
Understanding variations in dustbox use is absolutely
critical to the successful use of the method for ectoparasite
management in entire hen flocks. Dustbathing behaviour
consists of lying down and tossing litter onto and between
the feathers using a stereotypical series of scratching and
rubbing motions (van Liere, 1991). Dustbathing removes
excess lipids and maintains feather condition (Borchelt &
Duncan, 1974; Moyer et al., 2003). Dustbathing tends to occur
more frequently at midday and individual dustbathing sessions
average 27 min every 2 days (Vestergaard, 1982). Duncan
et al. (1998) demonstrated that domestic hens performed more
dustbathing under stimuli of heat and light, as well as when
hens in adjoining pens were dustbathing. Shimmura et al.
(2010) found 71–79% of test hens in all experiments performed
dustbathing in a dustbox, which is similar to our observations.
Dominant hens may take priority in the group’s use of the
resource (Shimmura et al., 2007). In our trials, certain hens
tended to use the dustbox repeatedly over time, whereas others
never did (data not shown), implying a possible hierarchy.
Several authors have noted that dustbathing effects on
ectoparasites have never previously been demonstrated experimentally
in wild or domestic birds (van Liere, 1992; Olsson
& Keeling, 2005; Clayton et al., 2010). The present study
shows dustbathing in naturally available substrates can suppress
ectoparasites. Kaolin, in particular, is one of the most
abundant soil constituents on earth. Silicaceous clays are
extremely common in soil surface strata as a consequence of
weathering (Singer & Munns, 2002). Given that birds greatly
prefer fine substrates for dustbathing (Olsson & Keeling, 2005),
and that rather pure deposits of kaolin or other clays are
so available, it is reasonable to assume birds dustbathe in
kaolin-like substrates and probably benefit from it in terms of
ectoparasite suppression. Diatomaceous earth and sulphur are
also natural materials and can cause ectoparasite suppression,
but areas of concentration are more restricted geographically.
Cage-free hens in our study had constant access to straw
litter material, which degraded into finer particles and mixed
with manure and feed. The proportion of very fine particles,
however, was minimal relative to that in the kaolin or DE and
sand mixtures supplied in the dustboxes. Only about 1% of
the litter particles were <250 μm in diameter, whereas clay is
defined by particle sizes of <2 μm (Singer & Munns, 2002).
The litter particles represented the finest particles available to
the cage-free hens. Nevertheless, the relatively coarse particles
in the litter were apparently insufficiently fine to impact on
ectoparasite suppression in a manner comparative with those
of the kaolin and DE used in the dustboxes. Dustbathing in
untreated litter appeared to have no significant impact on mite
or louse abundance when ectoparasite numbers were compared
with those on caged birds. Similarly, dustbathing in sand alone,
which is much coarser than kaolin or DE dusts, seemed to
have no impact on louse numbers in the studies by Hoffman
& Hogan (1967).
Certain criteria must be met in order for a behaviour such as
dustbathing to be considered adaptive (Hart, 1997). Firstly, the
parasite(s) must have detrimental effects on host fitness. Both
NFM and CBL have been shown to reduce egg production in
poultry (DeVaney, 1976; Mullens et al., 2009). Ectoparasites can also cause severe irritation to hens as well as personnel (Mullens et al., 2004b). Secondly, the behaviour must be
shown to reduce parasite numbers or severity. The present
study shows that dustbathing in a natural and widely available
substrate can effectively suppress ectoparasites, although it
would be desirable to test this with wild birds in a natural
setting. This distinctive behaviour is likely to represent an
evolutionary adaptation to suppress ectoparasites, as well as
to serve a feather maintenance function.
(Ornithonyssus sylviarum) and chicken body lice
(Menacanthus stramineus) on hens
C. D. MAR T I N and B. A. MUL L E NS
Department of Entomology, University of California Riverside, Riverside, CA, U.S.A.
Abstract. Hen housing (cage or cage-free) did not impact overall abundances
of northern fowl mites, Ornithonyssus sylviarum (Canestrini & Fanzago)
(Acari: Macronyssidae), or chicken body lice, Menacanthus stramineus (Nitzsch)
(Phthiraptera: Menoponidae). Cage-free hens received a dustbox with sand plus
diatomaceous earth (DE), kaolin clay or sulphur. Weekly use varied from none to
100% of hens; 73% of hens used the dustbox at least once. Ectoparasite populations
on dustbathing hens (users) were compared with those on non-user cage-free and caged
hens. All materials reduced ectoparasites on user hens by 80–100% after 1 week of
dustbox use. Diatomaceous earth and kaolin failed to reduce ectoparasites on non-user
hens, and ectoparasites on user hens recovered after dustbox removal. A sulphur dustbox
eliminated mites from all hens (including non-users) within 2–4 weeks. Residual
sulphur controlled mites until the end of the experiment (up to 9 weeks), even after
the dustbox was removed. Louse populations on hens using the sulphur dustbox were
reduced in 1–2 weeks. Residual sulphur effects were less evident in lice, but the use
of a sulphur dustbox by a higher proportion of hens extended louse control to all
hens. This is the first experimental study to show that bird dustbathing in naturally
and widely available dust materials (particularly kaolin) can suppress ectoparasites
and thus the behaviour is probably adaptive.
The northern fowl mite (NFM), Ornithonyssus sylviarum,
and the chicken body louse (CBL), Menacanthus stramineus,
are primary ectoparasites of poultry (DeVaney, 1978; Axtell
& Arends, 1990). Current control tactics consist primarily
of direct pesticide treatments to the host feathers and, in
some cases, treatments of the hen housing environment (i.e.
litter, nestboxes) to kill any ectoparasites that linger between
flocks. Unfortunately for producers, few synthetic chemicals
are available for mite control and resistance is problematic
(Mullens et al., 2004a). Further, although chemical control
and population dynamics of NFM and CBL have been
studied extensively in cage settings (Matthysse et al., 1974;
Arthur & Axtell, 1983; Mullens et al., 2009), scientific studies
examining mite and louse population dynamics and control in
hens held in cage-free conditions are lacking.
Animal welfare issues also are becoming increasingly important
in poultry systems (Rodenburg et al., 2008; Lay et al.,
2010). This has resulted in mandated changes in poultry
housing, such as the Welfare of Farmed Animals Regulations
Act of 2000 (http://www.opsi.gov.uk), and California’s
Proposition 2 (Standards for Confining Farm Animals)
(Sumner et al., 2008). In general, changes require moving
away from typical wire (‘battery’) cages, and providing hens
with greater space and/or altered or more complex cage
designs. Such changes could result in a potentially serious
economic disadvantage in states such as California relative
to other states lacking those requirements (Sumner et al.,
Pest control in the poultry industry must be tailored to the
different production systems (e.g. cages, enriched cages, cagefree,
free range). Notably, cage-free flocks cannot be treated
as easily as hens in cages, which can be treated using highpressure
chemical spraying from below. Even this method,
however, can result in incomplete coverage (Axtell, 1999).
Dustbathing behaviours have been described in chickens
(van Liere, 1991), wrens (Hendricks & Hendricks, 1995) and
quail (Borchelt et al., 1973), but few studies have experimentally
addressed the adaptive value of the behaviour. Literature
reviews have discussed the proven significance of dustbathing
for feather maintenance and have hypothesized that it reduces
ectoparasite numbers, but dustbathing effects on ectoparasites
have never actually been demonstrated experimentally in wild
or domestic birds (van Liere, 1992; Olsson & Keeling, 2005;
Clayton et al., 2010). Earlier studies tested pesticides or biological
agents in dustboxes for ectoparasite control, but individual
hen dustbox use and effects of untreated dust were not well
evaluated (Rodriguez & Riehl, 1957, 1960; Hoffman & Hogan,
1967; Hoffman & Gingrich, 1968).
The present study compared the population dynamics and
abundances of NFM and CBL in cage vs. cage-free settings. It
also explored dustbox use and the self-application of inorganic
dust materials as alternatives to the use of traditional synthetic
pesticides. Finally, the dustbox studies served as the first
experimental evaluation of the potential adaptive value of
dustbathing for ectoparasite suppression using available natural
substrates.
Materials and methods
Experiment design
Studies were conducted at the University of California (UC)
Agricultural Experiment Station next to the UC Riverside
campus, and procedures were approved through the UC
Riverside Institutional Animal Care and Use Committee.
Three trials were conducted between late autumn and late
spring (Trial 1: March–June 2009; Trial 2: October 2009 to
February 2010; Trial 3: March–June 2010). Beak-trimmed Isa-
Brown (Trials 1 and 3) or Hyline Brown (Trial 2) pullets were
obtained from a commercial operator. These breeds are popular
in cage and cage-free systems.
The research poultry houses (Fig. 1) measured 3.8 × 5.8 m,
were screened to exclude the entry of wild birds and
rodents, and equipped with roof sprinklers to maintain internal
temperatures below 33–35 ◦C. Hobo temperature recorders
(Onset Computer Corp., Bourne, MA, U.S.A.) recorded air
temperatures in each house. All hens had ad libitum access
to commercial lay mash in feed troughs and water was
continuously available. Lights inside the houses provided an
LD 16 : 8 h light cycle year-round, as is usual in commercial
production.
Twelve caged hens were held in each of two houses in
suspended wire cages. A rate of two hens per 31 × 31-cm
cage met U.S. United Egg Producer Standards (2010). Cagefree
hens were maintained in each of two houses, divided by
a plastic tarpaulin into north and south halves (3.8 × 2.9 m).
This allowed a dustbox treatment in one half of the house
to be compared with a no-dustbox treatment among cage-free
controls in the other half of the house (12 hens per treatment).
Cage-free houses contained eight nestboxes and perch space.
Each nestbox was about 0.1 m2 in area and was elevated about
0.8 m above the ground. Straw (2–4 cm deep) covered the
concrete floor At 19 weeks, 18 weeks and 20 weeks of age (in Trials 1–3,
respectively), each hen was infested with either 20–30 mites
(mostly adult females) or 20 lice (mostly adults, mixed sexes).
Mites were collected from a colony originally established in
2003. Mites were aspirated into capillary pipettes from source
hens and transferred to the vent area feathers (in an 8–9-cm
diameter zone extending from the base of the legs to the posterior
end of the cloaca) and skin of test hens, using a fine paint
brush. Lice (from a colony originally established in 2005) were
collected from source hens by inserting the hose from a carbon
dioxide (CO2) tank into the vent and abdominal feathers. This
lightly anaesthetized the lice, which were then shaken from the
abdominal feathers into a small bucket from which they were
transferred using a fine brush to the abdominal feathers of their
new hosts. One caged hen group and one cage-free hen group
were infested with NFM. Likewise, one caged hen group and
one cage-free hen group were infested with CBL.
All hens were identified as individuals by leg-banding one
of the two hens per cage and by using unique combinations
of coloured bands on the two legs of cage-free birds.
Ectoparasite counts were conducted weekly for 14 weeks
for NFM and 15 weeks for CBL. Inspections for NFM and
CBL were performed on different days to prevent crosscontamination.
Only the area near the vent was inspected
visually for NFM, using a scoring system (Arthur & Axtell,
1982) modified by Owen et al. (2009). Briefly, a score
of 0 = no mites seen, 1 = 1–10 mites, 2 = 11–50 mites,
3 = 51–100 mites, 4 = 101–500 mites, 5 = 501–1000 mites,
6 = 1001–10 000 mites, and 7 = >10 000 mites. A plus (+)
or minus (−) (except for a score of 7) was used to indicate a
population level in the upper or lower 20% of the respective
range.
In inspections for CBL, the chicken vent region was
examined as described for NFM. The area under each wing,
measuring approximately 8–10 cm in length and 5–7 cm in
width, consisting of the axilla and three feather partings down
the side of the body (separated by about 1.5–2.0 cm) was
also observed. Lice from the vent and under-wing areas were
summed.
Dustbox studies
Dustboxes (Fig. 2) had a 1.9-cm-thick plywood bottom, 2.5-
cm-thick wood sides and measured 65 × 65 × 20 cm. A 5-cmwide
wooden lip helped keep the particle mixtures in the box.
Washed playground sand (Premium Play Sand
; QuikreteCompanies, Inc., Atlanta, GA, U.S.A.) consisted of silica
particles of 0.5–1.5 mm in diameter. A 9 : 1 ratio by weight
of sand to dust [by volume: about 50 : 50 for diatomaceous
earth (DE) : sand and kaolin : sand, and about 25 : 75 for
sulphur : sand] was used in all three trials. The starting mixture
consisted of 10 800 g of sand and 1200 g of 100% DE
(Organic D/Earth
; Thomas Laboratories, Inc., Tolleson, AZ,U.S.A.), 95% kaolin (Surround WP
; Engelhard Corp., Iselin,NJ, U.S.A.) or 90% wettable sulphur (Yellow Jacket
; GeorgiaGulf Sulfur Corp., Valdosta, GA, U.S.A.), which filled the
dustbox to a depth of 3.5–5.0 cm. A small amount (0.16% by
weight) of fluorescent dust (FD; ‘arc yellow’ or ‘signal green’;
Day-Glo Color Corp., Cleveland, OH, U.S.A.) was added as
a dust marker (Hall et al., 1981) to estimate dustbox use by
individual hens. By shining a portable longwave ultraviolet
flashlight (WARD’S Natural Science, Rochester, NY, U.S.A.)
into the abdominal feathers under subdued ambient light, we
were able to assign a dustbathing score based on a combination
of the fluorescence level and overall visual ‘dustiness’ of
the hen. Scores were: 0 = no dust; 1 = trace dust (<25%
coverage of vent area, usually <10 individual FD particles);
2 = moderate dust (FD covering 25–50% of inspected vent
area), and 3 = high dust (FD covering 50–100% of inspected
vent area and particles often made airborne when parting
through feathers).
Each trial ran for 15 weeks; caged (no-dustbox) hens
were checked for ectoparasite numbers weekly. Cage-free
mite hens received a dustbox for weeks 4–8 [north end
(N)] and weeks 8–12 [south end (S)]. Cage-free louse hens
received a dustbox for weeks 5–9 (N) and 9–13 (S). This
provided experimental (dustbox treatment) replication with
two groups of hens for each ectoparasite. Other than those
4-week periods, the cage-free hens did not have access to
a dustbox. Thus, mites and lice on cage-free (N) hens were
given 4 weeks and 5 weeks, respectively, to reproduce before
hens were given access to a dustbox. Mites and lice on cagefree
(S) hens were given 8 weeks and 9 weeks, respectively,
to reproduce before hens were given a dustbox. Each cagefree
hen group was given a single dustbox for a 4-week
period, after which the box was removed. Dustboxes in
Trial 1 (DE) were replenished every other week with 50% of the starting material amounts; dustboxes in Trial 2 (kaolin)
were replenished weekly with 25% of the starting material
amounts. This replenishment was carried out to keep the
dustboxes effective because they appeared to lose the fine dust
material as the chickens scratched and kicked it out during
dustbathing. Trial 3 dustboxes were never replenished because
the effectiveness of the sulphur quickly became evident and
replenishment seemed unnecessary.
Data analysis
Statistical analyses were carried out using Minitab Version
14 (Minitab, Inc., State College, PA, U.S.A.). For parametric
analyses, the normality of each dataset was assessed using
the Ryan–Joiner test. One-way analyses of variance (anovas)
were used within a trial to generate weekly means for graphing and to compare weekly ectoparasite (NFM or CBL)
populations between hen groups. Housing effects (pre-dustbox)
data were also subjected to repeated-measures anova.
Regression slopes (abundance vs. time) were utilized as a
general indicator of population trends after dustbox removal
(mainly for DE and kaolin treatments). Slopes for the ‘during’
dustbox phase were based on a 5-week period, including the
ectoparasite level at the beginning of dustbox placement, and
those for each week until dustbox removal. Slopes used for the
‘after’ dustbox phase were based on periods of 3 weeks (cagefree,
S) to 5 weeks (cage-free, N), including the ectoparasite
level at the time of dustbox removal. This spanned the period
of 2–4 weeks post-dustbox removal. The effects of dustbox
use on ectoparasites were analysed by contrasting populations
on ‘users’ (FD scores of 2 or 3) with those on ‘non-users’
(FD scores of 0 or 1), employing two-sample t -tests. To
compare relative, short-term ectoparasiticidal effects of the
three materials, converted mite scores or louse numbers on
dustbathing hens (from week 0 to week 1 of dustbox use) were
compared with those on caged controls. Using the Henderson
& Tilton (1955) formula to calculate corrected percentage
control, the relative effects could then be determined by:corrected %
=
⎛
⎝1 −
n in Co before treatment
× n in T after treatment
n in Co after treatment
× n in T before treatment
⎞
⎠ × 100
where n = pest population, T = treated hens, and Co = control
hens.
In these analyses, mite indices were converted to approximate
‘actual’ values (0 = 0, 1− = 2, 1 = 5, 1+ = 8, 2− =
15, 2 = 30, 2+ = 45, 3− = 55, 3 = 75, 3+ = 95, 4− = 150,
4 = 300, 4+ = 450, 5− = 550, 5 = 750, 5+ = 950, 6− =
2000, 6 = 5000, 6+ = 8500, 7 = 20 000) to resolve dustbox
effects.
Results
Housing effects: northern fowl mites
Average weekly mite indices are illustrated in Fig. 3. To
isolate housing effects, these analyses reflect ectoparasite levels
prior to dustbox placement. Scores for NFM at equivalent
stages post-inoculation varied significantly between trials
(F = 13.41; d.f. = 2, 470; P <0.001). Therefore, trials were
examined individually.
Northern fowl mite population growth did not differ overall
between cage and cage-free treatments (weeks 1–
in any ofthe three trials (F ≤ 2.58; d.f. = 1, 145 or 1, 154; P ≥ 0.123).
Average weekly scores in the two housing types differed
occasionally (Fig. 3), but the variations were relatively minor
and not consistent. Differences between the cage-free (N)
treatment vs. the cage treatment were seen in Trials 1 and 2
(F <8.91; d.f. = 1, 66; P <0.03), but not in Trial 3 (F =
0.01; d.f. = 1, 66; P = 0.913). When weekly differences were
seen between the north and south cage-free flocks, the occurred in the hotter weather conditions of Trial 1. Degree
days (DD) above 25 ◦C were highest in Trial 1 (DD = 63.
and lower in Trials 2 and 3 (DD = 12.3 and DD = 7.3,
respectively). Both housing types allowed large numbers of
mites to develop on hens.
Housing effects: chicken body lice
Average louse numbers per hen prior to dustbox placement
differed significantly among the three trials (F = 25.32;
d.f. = 2, 540; P <0.001) (Fig. 4) and therefore trials were
examined separately. Louse abundance in cage vs. cagefree
(S) treatments did not differ significantly (weeks 1–9)
in any of the three trials (F ≤ 2.75; d.f. = 1, 172 or 1, 176;
P ≥ 0.112). Similarly, overall cage and cage-free (N) numbers
(weeks 1–5) did not differ significantly in any of the three
trials (F ≤ 1.44; d.f. = 1, 88; P ≥ 0.242). The rare weekly
differences were inconsistent. Both housing types allowed large
numbers of ectoparasites to develop on hens.
Dustbox studies: general trends
Figures 5 and 6 illustrate average on-host population trends
for NFM and CBL, respectively, in Trials 1–3 and depic three phases in cage-free treatments: (a) increasing ectoparasite
loads to high levels prior to dustbox placement; (b) ectoparasite
decline caused by hen dustbathing, and (c) general recovery
or stability of mite and louse populations after the removal
of DE and kaolin dustboxes. In general, standard errors of
mean ectoparasite numbers were higher when dustboxes were
present, reflecting differential dustbox use among hens.
Overall, mites and lice were reduced most effectively on
dustbathing hens, whereas non-user hens maintained moderate
to high levels of infestation (except for non-user hens in
houses with a sulphur dustbox). Figures 7 and 8 show average
ectoparasite levels on dustbox users compared with non-users
from week to week, and also indicate how many hens were
dustbox users in that week.
Over time, the straw on the cage-free floors degraded into
finer particles as the hens dustbathed, walked and scratched
in it, and straw particles became mixed with manure and
feed. As well as dustbathing in the boxes, hens would do
this in the litter before, during and after dustbox placement.
Based on the sieving of four random 1-liter samples at the
end of Trial 3, the particle size distribution [mean ± standard
deviation (SD)] was as follows: particles measuring >2.4 mm
accounted for 63 ± 16% of samples; particles of 0.6–2.4 mm
for 31 ± 10% of samples; particles of 0.25–0.6 mm for 5 ± 5%
of samples, and particles of <0.25 mm for 1 ± 1% of samples.
Notably, dustbathing in the litter apparently had no impact
on ectoparasite populations on cage-free hens; their numbers were similar to those on caged hens that were denied any
dustbathing opportunities. There were also obvious effects of
experimental dustbox use on ectoparasites when the boxes were
available (see below). These effects across time were clear
when contrasted with effects during periods when cage-free
hens could dustbathe only in litter.
Dustbox studies: diatomaceous earth
After 1 week of fresh dustbox access (week 4–5), average
mite counts (converted from NFM indices) on the six cagefree
(N) dustbathing (user) hens declined by 86% (75–100%
per hen) (Fig. 7A), whereas average mite counts on the six
cage-free (N) non-dustbathing hens increased by 30%. Mites
on cage-free, untreated hens (S) dropped by 61% and mites on
caged hens increased by 4% (Fig. 5A). User hen mite indices
were significantly lower than indices for non-user hens for
weeks 5, 6 and 8 (P ≤ 0.05) (Fig. 7A).
In a replication allowing fresh dustbox access with the
second hen group (week 8–9), mite numbers on 10 cagefree
(S) dustbathing hens declined by 79% (Fig. 7B), whereas
numbers on cage-free (S) non-user and caged hens dropped by
33% and 55%, respectively (Figs 7A and 5B, respectively).
Hens using the dustbox had significantly fewer mites than
non-user hens at weeks 11 and 12 (P ≤ 0.01) (Fig. 7B). Mite score regression slopes post-dustbox removal in Trial 1
did not differ significantly from zero (P ≥ 0.114). Mites on
caged hens during these periods also exhibited slight overall
negative regression slopes, but, again, these did not differ
significantly from zero (P ≥ 0.139).
After 1 week of dustbox access (week 5–6), average cagefree
(N) louse counts declined by 88% on the three dustbox
user hens and increased by 10% on non-users (Fig. 8A). In
the same period, louse populations on caged and untreated
cage-free (S) hens increased by an average of 50% and 40%,
respectively (Fig. 6A, C). Average louse counts on user hens
were significantly lower than on non-user hens (P ≤ 0.05)
(Fig. 8A) in every week of dustbox access.
In a replication with the second hen group (week 9–10),
average louse counts on nine of 11 cage-free (S) dustbathing
hens declined by 96%, whereas average counts on non-users
declined by 26% (Fig. 8B), and counts on caged controls
declined by 14% (Fig. 6A). No significant differences in louse
numbers were detectable between user hens and the very few
non-user hens in weeks 10–13 (Fig. 8B).
Regressions of louse numbers over time in the weeks
following DE dustbox removal produced positive slopes in
both treatments, although neither differed significantly from
zero (P ≥ 0.251) (Fig. 6B, C). Similarly, lice on caged hens exhibited no significant slope deviation during this recovery
period.
Dustbox studies: kaolin
After 1 week of fresh dustbox access (week 4–5), average
mite abundance on five of 12 cage-free (N) dustbathing hens
declined by 95%. Mite numbers on seven of 12 cage-free (N)
dustbox non-user hens increased by 10% (Fig. 7C). Mites on
untreated caged and cage-free (S) control groups increased by
230% and 370%, respectively (Fig. 5A, C).
In a replication with the second hen group (week 8–9), mite
numbers on six of 12 cage-free (S) dustbathing hens declined
by 90% after treatment and those on non-users remained stable
(Fig. 7D). Numbers on caged controls and cage-free (S) nonusers
increased by 80% and 50%, respectively (Fig. 6A, C).
Numbers of mites recovered upon dustbox removal in
Trial 2, as indicated by significantly positive regression slopes.
cage-free (N) scores exhibited a slope significantly greater than
zero (P = 0.003) and increased until mite populations reached
levels approximate to those on caged birds.
Average louse numbers declined by 88% after 1 week of
dustbox access (week 5–6) on six of 12 cage-free (N) dustbox
users but increased by 90% on non-user hens (Fig. 8C).
Meanwhile, louse abundance on caged and cage-free (S)
control groups increased by 70% and 110%, respectively
(Fig. 6A, C).
In a replication with the second hen group, the cage-free (S)
hens in this trial did not accept the dustbox in the first week
(week 9–10) for unknown reasons. By the second week, two
of 12 hens used the resource. Louse numbers on these two
birds were reduced by 73% and 98% (Fig. 8D).
Louse population regression slopes were positive for the
period post-kaolin dustbox removal in both cage-free treatments.
The slope for cage-free (N) louse trends in Trial 2
(post-dustbox) was significantly positive (P = 0.005).
Dustbox studies: sulphur
After dustbox access (week 4–5), mite numbers on three
of the 12 cage-free (N) dustbathing hens declined by 99.9%
(Fig. 7E). Interestingly, mite numbers on nine of 12 cagefree
(N) non-dustbathing hens also declined by 99%. Mite
numbers on caged and cage-free (S) control groups dropped
by 12% and 35%, respectively (Fig. 5A, C).
In a replication with the second hen group (week 8–9),
mite numbers on three cage-free (S) dustbathing hens declined
by 99.9% (Fig. 7F), whereas numbers on caged and cagefree
(S) non-user hens dropped by 19% and 29%, respectively
(Fig. 5A, C). Dustbathing in sulphur was 100% effective in
reducing mites to low levels (scores of <2) after 1 week of
access, and all visible mites were eliminated in a 2–4-week
period (Fig. 5B, C).
Mites did not recover from sulphur dustbox treatments
up to 6 weeks after dustbox removal (Fig. 5B, C), although
relatively few (≤50%) of the hens in each group used the
dustbox on a weekly basis.
After 1 week of dustbox access (week 5–6), lice on the
three cage-free (N) user hens dropped by 100% (Fig. 8E).
Interestingly, louse numbers dropped by 88% on non-user
hens, whereas louse numbers on caged controls increased by
40% (Fig. 6A).
In a replication with the second hen group, lice on
seven cage-free (S) user hens (week 9–10) dropped by 98% (Fig. 8F). Louse numbers on non-user hens and
caged controls dropped by 90% and 47%, respectively
(Fig. 6A).
Louse numbers recovered quickly following sulphur dustbox
treatment when fewer hens used the dustbox (Fig. 6B), but
no recovery was observed up to 2 weeks following dustbox
removal after heavier hen use (Fig. 6C). By comparing ‘before’ and ‘after’ ectoparasite levels on
cage-free hens (user hens only) with ‘before’ and ‘after’
levels on caged hen controls (week 0–1 of dustbox use), the
Henderson & Tilton (1955) formula could be used to calculate
corrected percentage short-term control for DE, kaolin and
sulphur treatments (Table 1). Sulphur was the most active
material. Kaolin effects tended to be more active in the short
term than those of DE, particularly for mites. However, DE was
replenished every 2 weeks, whereas kaolin was replenished
weekly.
Discussion
Housing
Ectoparasite abundances on poultry were not substantially
or consistently affected by housing (cage vs. cage-free). Hens
in both cage and cage-free settings were equally capable of
supporting high ectoparasite loads when untreated with dustboxes
(Figs 3 and 4). Using only caged hens, Hall et al.
(1978), Arthur & Axtell (1983) and Mullens et al. (2000)
observed fewer mites under more crowded cage conditions.
In the present study, mite populations on cage-free hens occasionally
tended to be higher early in infestation than those on
hens in cages, but large or consistent ‘density’ effects on mites
were not seen. Mullens et al. (2010) showed that hens held
in pairs per cage supported about twice as many lice as hens
held singly in cages, presumably because crowded hens were
less efficient in grooming. In the present study, no significant
housing effects on CBL abundance emerged in any trial.
Dustboxes
Hen dustbathing in DE and kaolin dustboxes had very
clear, immediate, negative impacts on mite and louse numbers.
These effects were temporary, however, and depended on
the individual hen’s use of the dustbox. Dustboxes had beensuggested for managing mite and louse infestations on poultry farms at least by the early 1900s (Banks, 1907; Pierce &
Webster, 1909). The few existing dustbox studies evaluated
the effects of synthetic insecticides, rather than documenting
untreated substrate effects on ectoparasites (Rodriguez &
Riehl, 1957, 1960; Hoffman & Hogan, 1967). Hoffman &
Hogan (1967) attributed a lack of louse control to nonacceptance
or monopolization of an insecticidal dustbox, but
did not document dustbox use. Hoffman & Gingrich (1968)
conducted a similar study testing hand vs. dustbox applications
of Bacillus thuringiensis Berliner. They demonstrated little to
no control of lice in dustbox treatments compared with direct
hand applications, but again did not document dustbox use.
Diatomaceous earth and kaolin
Dry formulations of DE and kaolin are relatively nontoxic
to humans, although they do pose some inhalation risks
(Lynch & McIver, 1954; Merget et al., 2002). The presumed
mode of action of these materials is arthropod death by
desiccation, which is achieved either by abrading the cuticle
or absorbing lipids from the cuticle that are important for
managing water loss (Wigglesworth, 1945; Ebeling, 1971).
Ebeling (1973) described kaolin clay as a slightly abrasive
material and DE as both abrasive and sorptive. Cook et al.
(2008) found no actual evidence of abrasion of flour mites
[Acarus siro (Sarcoptiformes: Acaridae)] after exposure to
three DE products for 3–72 h. However, significant amounts
of tridecane, a cuticular lipid thought to be an important
waterproofing agent, were detected on all of the DE products
rinsed off the mites. This suggests the main action of the
dusts, at least on mites, may be through absorption, rather than
abrasion.
Inert dusts, such as DE, are used to treat poultry houses for
Dermanyssus (Acari: Dermanyssidae) in Europe (Mul et al.,
2009; Harrington et al., 2011). Dawson (2004) suppressed fleas
and Protocalliphora (Diptera: Calliphoridae) in tree swallow
nests using DE. Kilpinen & Steenberg (2009) tested DE and
kaolin for control of D. gallinae (De Geer) and found that both
killed mites, but the efficacy of each material decreased with
increasing (75–85%) relative humidity. Based on 24-h miteweight loss studies, kaolin clay resulted in 11.9% loss and DE
exposure in 18.2% loss, compared with 9.5% weight loss in
a control group (Kilpinen & Steenberg, 2009). Despite blood
engorgement by this mite, the dusts may cause water loss faster
than it can be replenished.
Sulphur
Sulphur is a natural component of the environment; it is
generally considered to be safe and is currently the active
ingredient in nearly 300 pesticides (http://www.epa.gov). In
small concentrations (mixed with petroleum jelly), it is used
as a topical treatment for control of scabies mites on humans
(Pruksachatkunakorn et al., 2002). As a dust, however, it is
considered slightly toxic and slightly irritating in terms of
inhalation and eye exposure, and should be kept away from
sparks and flames because it has some ignition potential.
Sulphur has probably been used for mite and louse control
in poultry longer than any other material. Sulphur kills by
direct contact and is thought to have fumigant action at
temperatures >21 ◦C (Ware, 1993), which may be especially
effective within the feather layer. Banks (1907) recognized that
sulphur and lime mixtures could be scattered in a poultry house
for control of mites (D. gallinae) and lice and commented on
the fumigant power of sulphur against lice. Creighton et al.
(1943) tested sulphur for control of poultry lice on White
Leghorns as a feed supplement, a direct dust application and
a soil treatment. They reported that consumption by chickens
was ineffective for control, but that direct and soil treatments achieved excellent control within 1–2 weeks, and that residual
effects were poor when test chickens were later exposed to
heavily infested chickens and were re-infested (Creighton et al.
1943). Furman (1952) showed hand-dusting was effective for
NFM control on hens, and Foulk & Matthysse (1963) achieved
success when dusting caged hens for NFM.
Sulphur was clearly the most potent material tested in our
dustbox studies. User hen groups were most affected, but rather
quickly and over time sulphur had effects on ectoparasites
even on non-user hens. Mites were controlled for ≥9 weeks,
although fewer than 50% of the chickens might use the dustbox
in a given week. In a hen group, lice recovered from sulphur
dustbox treatments when few hens were using the dustbox, but
were severely suppressed for ≤3 weeks on all chickens if more
hens used it. This implies that dustbathing hens were largely
responsible for dispersing sulphur in the environment.
Dustbathing and dustbox use
The most significant factor influencing the value of a dustbox
treatment was the variation in the individual’s use of the box.
About 73% of all hens that had access to a dustbox used
it at some point. Some used it more than others, and some
did not use it at all. In previous dustbox studies, dustboxes
were considered unreliable as a control method for mites and
lice because of differences in dustbox usage among individual
hens (Bishopp & Wood, 1917, 1931; Hoffman & Hogan, 1967;
Hoffman & Gingrich, 1968).
Understanding variations in dustbox use is absolutely
critical to the successful use of the method for ectoparasite
management in entire hen flocks. Dustbathing behaviour
consists of lying down and tossing litter onto and between
the feathers using a stereotypical series of scratching and
rubbing motions (van Liere, 1991). Dustbathing removes
excess lipids and maintains feather condition (Borchelt &
Duncan, 1974; Moyer et al., 2003). Dustbathing tends to occur
more frequently at midday and individual dustbathing sessions
average 27 min every 2 days (Vestergaard, 1982). Duncan
et al. (1998) demonstrated that domestic hens performed more
dustbathing under stimuli of heat and light, as well as when
hens in adjoining pens were dustbathing. Shimmura et al.
(2010) found 71–79% of test hens in all experiments performed
dustbathing in a dustbox, which is similar to our observations.
Dominant hens may take priority in the group’s use of the
resource (Shimmura et al., 2007). In our trials, certain hens
tended to use the dustbox repeatedly over time, whereas others
never did (data not shown), implying a possible hierarchy.
Several authors have noted that dustbathing effects on
ectoparasites have never previously been demonstrated experimentally
in wild or domestic birds (van Liere, 1992; Olsson
& Keeling, 2005; Clayton et al., 2010). The present study
shows dustbathing in naturally available substrates can suppress
ectoparasites. Kaolin, in particular, is one of the most
abundant soil constituents on earth. Silicaceous clays are
extremely common in soil surface strata as a consequence of
weathering (Singer & Munns, 2002). Given that birds greatly
prefer fine substrates for dustbathing (Olsson & Keeling, 2005),
and that rather pure deposits of kaolin or other clays are
so available, it is reasonable to assume birds dustbathe in
kaolin-like substrates and probably benefit from it in terms of
ectoparasite suppression. Diatomaceous earth and sulphur are
also natural materials and can cause ectoparasite suppression,
but areas of concentration are more restricted geographically.
Cage-free hens in our study had constant access to straw
litter material, which degraded into finer particles and mixed
with manure and feed. The proportion of very fine particles,
however, was minimal relative to that in the kaolin or DE and
sand mixtures supplied in the dustboxes. Only about 1% of
the litter particles were <250 μm in diameter, whereas clay is
defined by particle sizes of <2 μm (Singer & Munns, 2002).
The litter particles represented the finest particles available to
the cage-free hens. Nevertheless, the relatively coarse particles
in the litter were apparently insufficiently fine to impact on
ectoparasite suppression in a manner comparative with those
of the kaolin and DE used in the dustboxes. Dustbathing in
untreated litter appeared to have no significant impact on mite
or louse abundance when ectoparasite numbers were compared
with those on caged birds. Similarly, dustbathing in sand alone,
which is much coarser than kaolin or DE dusts, seemed to
have no impact on louse numbers in the studies by Hoffman
& Hogan (1967).
Certain criteria must be met in order for a behaviour such as
dustbathing to be considered adaptive (Hart, 1997). Firstly, the
parasite(s) must have detrimental effects on host fitness. Both
NFM and CBL have been shown to reduce egg production in
poultry (DeVaney, 1976; Mullens et al., 2009). Ectoparasites can also cause severe irritation to hens as well as personnel (Mullens et al., 2004b). Secondly, the behaviour must be
shown to reduce parasite numbers or severity. The present
study shows that dustbathing in a natural and widely available
substrate can effectively suppress ectoparasites, although it
would be desirable to test this with wild birds in a natural
setting. This distinctive behaviour is likely to represent an
evolutionary adaptation to suppress ectoparasites, as well as
to serve a feather maintenance function.
. What I would really like to see is a shortened version with just the results, what was obtained to get the results, in a short, short manner. I would love to try to get this stuff, but too much information that is just beyond what I want to do. I bet it is a very good read, and one day, maybe I will finish, only got about 1/3 the way through and gave up....have a wonderful day, CynthiaM.
Have a wonderful day, CynthiaM.
. have changed my clothes twice today and still feel crawlie!! grrrrrrrrrrrrr. might also have mites. need to do more thorough examination but lice for sure. I don't mean to open the whole topic up about what to use. Just want to know if anyone can report back about their experience with the sulfur, is anyone using it and has it been effective? I wonder about lice and the litter in the coop, do I have to clean the whole coop out? I was trying for the deep litter technique and so far like it. Any thoughts? It has been over a month since this post was started, wondering what the story is so far. 
. To which I shakily said "it's er . . . er it's for my chickens . . .?" She said "Oh" and left me hanging. So I worked a little and said I'm looking at different options and have friends who have show birds who use this stuff ........ well eventually she said you'd probably want the one for cats and it is $43 for three months worth for one cat, three tubes in the packet. We talked about how you'd want only a little drop. I agreed. 
So today I'm going to go check out my feed store and some other stores to maybe and get a cheap Rubbermaid container and let the girls go to town.