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Assessing reproductive profiles in female brown mouse lemurs (Microcebus rufus) from Ranomafana National Park southeast Madagascar using fecal hormone analysis.

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American Journal of Primatology 71:439–446 (2009)
Assessing Reproductive Profiles in Female Brown Mouse Lemurs (Microcebus
rufus) From Ranomafana National Park, Southeast Madagascar, Using Fecal
Hormone Analysis
Department of Anthropology, University of Massachusetts, Amherst, Massachusetts
Department of Psychology, University of Massachusetts, Amherst, Massachusetts
Studies on reproductive endocrinology in wild primate populations have greatly increased in the last
decades owing to the development of noninvasive techniques that can be applied under field conditions.
However, small-bodied nocturnal species are not well represented on the long list of primates surveyed
in the wild, and reproductive inferences regarding these animals in their natural habitats have not
benefited from direct observations of hormonal changes. We collected fecal samples from female brown
mouse lemurs (Microcebus rufus) in a southeastern rainforest of Madagascar in order to determine
whether or not fecally excreted steroid levels show a consistent pattern of change during the
reproductive season and are a useful complement to reproductive observations in wild-trapped
individuals. Initial data show variation in reproductive hormone levels before and after estrus and
estimated day of parturition. Elevated levels of excreted estradiol (E2) were observed around the time of
estrus, whereas high levels of fecal progesterone (P) were seen during later stages of pregnancy and
around parturition. A more complete picture of reproductive profiles in female mouse lemurs, and how
they may change over the life span, can be obtained if hormone analyses are used to supplement field
observations. Am. J. Primatol. 71:439–446, 2009.
r 2009 Wiley-Liss, Inc.
Key words: Microcebus rufus; Ranomafana National Park; estradiol; progesterone; fecal analysis
Noninvasive endocrine analyses of ovarian function have been applied successfully to a variety of
captive as well as wild primate populations. Ever
since the pioneering work by Risler et al. [1987] and
Wasser et al. [1988], different methodologies have
been explored and validated under captive conditions, where the reproductive status of individuals
can be regularly monitored and the relative efficacy
of different types of samples (e.g. urine, blood, feces)
can be assessed. The consensus that fecal steroids are
informative vis-à-vis reproductive function has
greatly influenced the field of primate endocrinology,
and it has facilitated the analyses of endangered
species for which only noninvasive techniques can be
applied [Strier & Ziegler, 1997]. To date, fecal
endocrine profiles that employ a single or a combination of reproductive hormones have been reported
for many primate groups. These include sifakas, red
fronted lemurs and mongoose lemurs among lemuroids [Brockman & Whitten, 1996; Brockman et al.,
1995; Curtis et al., 2000; Ostner & Heistermann,
2003]; tamarins, marmosets, muriquis and saki
monkeys among ceboids [Heistermann et al., 1993;
Shideler et al., 1994; Strier et al., 2003; Ziegler et al.,
1996]; a variety of macaques and langurs among
r 2009 Wiley-Liss, Inc.
cercopithecoids [Bardi et al., 2003; Heistermann
et al., 1995; Shideler et al., 1993]; and bonobos and
gorillas among hominoids [Atsalis & Margulis, 2006;
Jurke et al., 2002; Miyamoto et al., 2001].
Some groups, however, are poorly represented or
entirely missing from the list. In particular, there is
a paucity of data on small-bodied nocturnal lemurs in
the wild. This bias toward larger diurnal primates
may be in part logistical. It is difficult to collect fecal
samples during focal animal observations at night
when the animals are active; thus, feces are most
easily gathered when individuals are captured in
traps. Frequent capture and recapture of the same
individual may be required to obtain sufficient
samples to reconstruct its endocrine profile, a task
Contract grant sponsors: Rufford Foundation; MMBF/CI Primate Action Fund; Institute of Biotechnology; NSF; Contract
grant number: BCS-0721233.
Correspondence to: Marina B. Blanco, Department of Anthropology, 240 Hicks Way, University of Massachusetts, Amherst,
MA 01003. E-mail:
Received 15 March 2008; revised 8 December 2008; revision
accepted 18 January 2009
DOI 10.1002/ajp.20672
Published online 10 February 2009 in Wiley InterScience (www.
440 / Blanco and Meyer
that cannot be guaranteed (i.e. other individuals may
enter the same traps at any given night) and may
indeed disrupt reproductive schedules if the same
individual enter the same traps every single day.
Our study species, the brown mouse lemur,
Microcebus rufus, is a small-bodied (40 g) nocturnal
seasonal breeder that inhabits the eastern rainforests of Madagascar. Although brown mouse
lemurs as well as gray mouse lemurs (M. murinus),
a western species, have been kept in captivity for
decades [Glatston, 1979; Perret, 1986; Wrogemann &
Zimmermann, 2001], little has been published on
their endocrine profiles. Furthermore, there is
currently a lack of consensus regarding the utility
of fecal analysis and even less understanding of
individual variability in mouse lemur steroid profiles,
and particularly how such profiles may change, over
their life span.
Glatston [1979] conducted preliminary hormone
analysis in captive female mouse lemurs in an
attempt to detect the early stages of pregnancy, but
no urinary estrogen peaks were detected during
estrus or later. Further studies on reproductive
hormones have shown mixed results. In 1986, Perret
reported consistent plasma progesterone (P) levels in
consecutive estrous cycles of captive M. murinus
females. Variation in hormonal levels among individuals was attributed to different social conditions. A
subsequent study of the same species found, but
could not explain, high intra- and inter-individual
variation in serum P [Buesching et al., 1998]. The
latter study also attempted to use urinary and fecal
estrogen and P metabolites to monitor reproductive
function noninvasively; however, no clear pattern of
change in steroid levels within the reproductive cycle
was found. Finally, Perret [2005] showed a clear
progressive increase of urinary estradiol (E2) 10
days before estrus, a peak of excretion at ovulation,
and a consequent decrease to baseline levels within 4
days, although she reported significant variation
among individual females, and she did not test fecal
Collecting serum is highly invasive and collecting urine from wild-caught animals is logistically
difficult given the problems posed by preservation
and storage of urine under field conditions. Preservation of dried-oven fecal samples is easier, and it
seems possible that prior apparent negative results
may be owing to low assay sensitivity or hormonal
variability over the life span. This in itself may be of
biological interest. Because so little is known about
fecal steroids in mouse lemurs even in captivity,
sampling fecal steroids should not be abandoned
without more rigorous testing. We now report data
obtained from live-trapped mouse lemurs at Ranomafana National Park, a southeastern rain forest in
Madagascar. Our aims are to (1) determine whether
or not a clear pattern of fecal E2 and P excretion
exists in female brown mouse lemurs during the
Am. J. Primatol.
reproductive season; (2) document the efficacy of
minimally invasive fecal hormonal analysis as a
reliable marker of ovulation, parturition and early
lactation, within the context of overall individual
variability and (3) explore factors (such as age) that
might help us to understand the pattern of variation
among individuals. This study represents the first
report on fecal steroid analysis as a tool to monitor
reproductive function in wild mouse lemurs.
This research, conducted under permission of
institutional and governmental agencies that regulate animal research in Madagascar, adhered to the
American Society of Primatologists Principles for the
Ethical Treatment of Nonhuman Primates. Research
protocols complied with those approved by the
University of Massachusetts—Amherst Animal Care
and Use Committee.
Study Area and Field Observations
We conducted field work in the Talatakely Trail
System at Ranomafana National Park, a montane
rainforest in southeast Madagascar (471180 –471370 E
and 211020 –211250 S). We employed capture/mark/
recapture techniques in October–December 2005,
October 2006–January 2007 and October 2007 as a
part of a long-term monitoring of mouse lemur
populations in this study area (9 ha). A maximum
of 50 Sherman traps at intervals of ca. 25 m were
baited with a piece of fresh banana and set up daily
between 16:00 and 17:00 hr along preexisting trails
(similar areas have been surveyed over the years to
maximize recapture rates) on consecutive nights
(trapping nights: 69 in 2005; 54 in 2006, 21 in
2007). Traps were checked between 19:30 and
21:00 hr, and trapped animals were brought to the
ValBio research station where individuals were
identified or marked with Avid microchips, weighed
and handled to observe their reproductive status.
Trapped Microcebus were released between 23:00
and 1:00 hr on the same night of capture to minimize
disturbance of their reproductive schedules.
Reproductive status of females was determined by
inspecting vaginal morphology; i.e. diestrous condition
was characterized by sealed and swollen vaginas;
proestrous, estrous and metestrous conditions were
determined during vaginal openings through the
inspection of cytological features at 40 magnification from vaginal smears. Physiological estrus was
determined by the presence of sperm (decapitated
heads and/or tails), vaginal plugs or by the inspection
of cytological features from the smears following
descriptions for mouse lemurs available in the
literature [e.g. Blanco, 2008; Wrogemann & Zimmermann, 2001]; additional observations such as nipple
development and body mass were used to generally
distinguish the later stages of pregnancy and early
Fecal Excreted Steroids in Microcebus rufus / 441
lactation. More information about nipple development
and body mass changes during gestation is provided in
Blanco [2008]. Gestation length could be established
with confidence in three cases (‘‘I’’, ‘‘J’’ and ‘‘K’’ in
2005) giving a period of 57 days [Blanco, 2008].
These estimations agree with published records from
captivity [Wrogemann & Zimmermann, 2001]; the
date of parturition of the remaining females was then
estimated by adding 57 days to the date of estrus. We
could preliminarily classify the female population into
two broad age categories: young (1–2 years old) vs.
adults (Z3 years old) based on the pattern of dental
wear and body mass. Zodhy et al. [personal communication] have developed an age scale based on digital
measurements from high-quality dental molds
from frequently captured individuals. In addition,
1-year-old individuals tend to be lighter than older
females at the beginning of the reproductive season
[personal observation].
We collected fecal samples from animals that
defecated when handled. If fresh feces were not
available, then we recovered samples from inside the
Sherman traps where the individual females had
remained for a period no longer than 3 hr. Feces were
weighed with a digital balance (accurate to 0.01 g),
wrapped in aluminum foil, and oven-dried for about
2–3 days at 501C. Fecal pellets were then stored in
plastic bags at room temperature until the end of the
field season (3.5-month period). Once at the University of Massachusetts—Amherst, samples were
kept frozen at 201C until assayed. Except for
female ‘‘V’’ (for which samples collected in 2005
were assayed in 2007), all fecal samples were
analyzed within 6 months after being brought back
from the field.
We used SPSS 15.0 to calculate statistical
significance (a 5 0.05) in parametric (Independent
T-tests, Levene’s Tests for Equality of Variance) and
nonparametric tests (Sign Test). One-tailed tests of
significance were used to test specific directional
hypotheses—for example, that E2 should decrease
following ovulation (H0 5 there is no change in E2
over ovulation, or E2 increases following ovulation).
Fecal Sample Preparation
The protocol that we employed was modified
from that of Khan et al. [2002]. Fecal pellets were
ground with a Mixer Mill MM200 milling machine
(Retsch Inc., Newtown, PA). This method produces a
finer powder than would be obtained by hand
grinding, which should improve hormone extraction
from small samples. After milling, the samples were
weighed and transferred to 2 ml microcentrifuge
tubes containing an extraction solution consisting
of 1.5 ml 90% methanol and 50 ml of [3H]E2 (approximately 75,000 CPM; 83.0 Ci/mmol, Amersham Biosciences, PA) in ethanol as an internal standard. For
later determination of steroid recovery, 300 ml of the
same extraction solution were added to 10 ml of
scintillation cocktail (ScintisafeTMSX 23–5, Fisher
Scientific, PA) in duplicate and radioactivity was
measured in a Packard 1900CA scintillation counter.
Samples were extracted overnight at room temperature using a rotator. The following morning, samples
were centrifuged for 10 min (speed 5, Marathon
Micro A, Fisher Scientific) and 800 ml of the supernatant was transferred into a clean glass tube and
dried under nitrogen gas. Samples were reconstituted
in 400 ml of 30% methanol and vortexed intensively
before running the solid-phase extraction.
Fecal steroids were extracted using solid-phase
Oasis cartridges (3cc/60 mg HLB, Waters, Milford,
MA). Cartridges were conditioned with 1.0 ml of
100% methanol followed by 1.0 ml of deionized
water. Each cartridge was loaded with 300 ml of each
sample and rinsed with 1.0 ml of 20% methanol.
Steroids were eluted from the columns with 2.0 ml of
100% methanol [Khan et al., 2002]. Samples were
stored at 201C until assayed.
Steroid Radioimmunoassays (RIAs)
Before conducting steroid RIAs, samples were
dried under nitrogen gas and reconstituted with
300 ml of 30% methanol. One hundred microliter of
each sample were added to 10 ml of the scintillation
cocktail and radioactivity was measured to determine steroid recovery. For the E2 assay, 15 ml of each
sample was diluted in 1.485 ml of the diluent
provided in the RIA kit before being assayed. To
measure fecal E2 we employed the Pantex 125I
Estradiol kit (catalog ]047, Santa Barbara, CA) and
followed the manufacturer’s instructions. Crossreactivity provided by the manufacturer is 1.4% for
a-estradiol and less than 0.02% for other steroids.
Our estimated intra-assay Coefficient of Variation
(CV) for E2 was 11%. Briefly, serial dilutions of a
calibrated standard and the fecal extracts at a
volume of 500 ml were assayed in duplicate. One
hundred microliter of tracer was added to each tube,
followed by 100 ml of the first antiserum. After 30 min
of incubation at 371C, 500 ml of the second antiserum
was added to the samples and the standards. Tubes
were vortexed, left at room temperature for 10 min,
and then centrifuged at 2400 rpm for 15 min. Supernatants were discarded immediately and radioactivity of precipitates was counted using a Packard
gamma counter (CobraTM II, Packard Instrument
Company, CI). To assess levels of fecal P, we used the
Pantex 125I Direct Progesterone kit (catalog ]137,
Santa Barbara, CA), which is highly specific for P
with cross-reactivity of 0.5% for hydroxyprogesterone, 0.1% for androsterone and values below 0.1% for
other steroids. Intra-assay CV for P was 10.9%. For
the P assay, 50 ml of each sample was diluted in 450 ml
of diluent. The standards and the samples (100 ml)
were prepared in duplicate and 500 ml of tracer were
Am. J. Primatol.
442 / Blanco and Meyer
added to the tubes, followed by 100 ml of the first
antiserum. Tubes were vortexed and incubated at 371C
for 1 hr. Then 500 ml of the second antiserum was
added and tubes were left at room temperature for
10 min. Finally, the tubes were centrifuged at 2400 rpm
for 15 min, supernatants were immediately discarded,
and precipitate radioactivity was counted as before.
A total of 47 individual females were captured
during the reproductive seasons of 2005, 2006 and
2007. We determined physiological estrus from
vaginal smears (n 5 17) and estimated dates of
parturition for 10 individual females trapped during
the study period. One hundred twenty fecal samples
were assayed for excreted E2 and P. Females ‘‘V’’,
‘‘Ke’’ and ‘‘S’’ contributed fecal samples for two
consecutive years and ‘‘I’’ and ‘‘J’’ for all three
seasons. Hormone levels, expressed as pg/mg for E2
and ng/mg for P, were corrected for steroid recovery
in each sample, which averaged 73% in samples
assayed in 2008, and 76 and 77% in samples analyzed
in 2007 and 2006, respectively. In all, seven females
were sampled around the time of estrus for E2 (for a
total of 11 cases) and six females were sampled for P
during the same period (for a total of ten cases).
Around the estimated day of parturition, samples
from nine females were available for E2 (for a total of
18 cases) and P (for a total of 16 cases).
In nine of 11 cases, E2 levels were higher before
or at estrus (day 10 to day 0) than during the 5-day
period after estrus (day11 to day15). This difference
was statistically significant (Sign Test, one-tailed,
P 5 0.03, Fig. 1). There was substantial intra- and
inter-individual variation in E2, as illustrated by
maximum pre- and post-estrus E2 levels in Table I.
Variation was significantly higher in young than in
adult individuals (Table II), although the smaller
number of young animals may have contributed to
this difference. In six of ten cases, P levels increased
after estrus (day11 to 15) but generally remained
relatively low. The change from pre- to post-estrus in
the sample was not statistically significant (Sign
Test, one-tailed, P 5 0.37, Fig. 2). Figure 3 illustrates
the hormonal profile of individual ‘‘I’’ in 2006, which
is consistent with changes in vaginal morphology
around the time of estrus.
Owing to a lack of certainty concerning the exact
day of parturition for some of the pregnant females,
we designated the prepartum period to be 10 to 0
(71) days. E2 levels tended to increase during this
period followed by a decrease after parturition.
Although the post-partum pattern is clearly shown
in Figure 4, no significant differences were found,
possibly owing to small sample sizes (particularly the
few number of animals for which samples were
available for all three periods) as well as the high
intra- and inter-individual variation (Table IIIa).
Similarly, P levels were higher during the later
stages of pregnancy than during the weeks following
parturition (Fig. 5; Table IIIb).
Fig. 1. E2 levels before (10 to 0 days) and after estrus (11 to 15
days). Each symbol represents an individual’s mean values
within that period. One-tailed Sign Test, P 5 0.03.
TABLE I. Maximum E2 Values Before and After Estrus for Young (1–2-year-old) and Adult (Z3-year-old)
K 05
S 06
S 07
V 06
A 07
I 05
I 06
I 07
J 07
Ke 06
Ke 07
Est. age
Days from estrus
Days after estrus
Females are identified by initials and year of capture.
Am. J. Primatol.
Fecal Excreted Steroids in Microcebus rufus / 443
TABLE II. Mean E2 Values Prior to (10 to 0 days)
and After Estrus (11 to 15 days) for Young and Adult
Young (n 5 4)
Adults (n 5 7)
F 5 22.9, P 5 0.001.
F 5 140, Po0.001 by Levene’s Test for equality of variance.
Each mean is the average of the mean values for each of the individuals in
the sample.
Fig. 4. E2 levels before and after estimated parturition (7 1day).
Means were used for individual females with multiple values
within each period.
TABLE III. Descriptive Statistics for (a) E2 and (b) P
Before and After the Estimated day of Parturition
(71 day)
Time period
(a) Estradiol (E2)
[–10 to 10]1
[12 to 114]2
[115 to 131]3
Fig. 2. P levels before (10 to 0 days) and after estrus (11 to 15
days). Each symbol represents an individual’s mean values
within that period. One-tailed Sign Test, P 5 0.38.
Fig. 3. Hormone profile of female I 06 around the time of estrus.
E2 peaked before vaginal opening and P began to rise at the end
of the first week of pregnancy. Reproductive observations: 7: no
vaginal opening; 4: swollen vagina; 2: very swollen vagina;
0: vaginal plug; 15: vaginal opening, but in the process of
(b) Progesterone (P)b
[10 to 10]1
[12 to 114]2
[115 to 131]3
Independent sample t-test for equivalence of means: 1,2t 5 1.7, df 5 13,
P 5 0.1; 2,3t 5 0.3, df 5 9; P 5 0.8; 1,3t 5 1.2, df 5 12, P 5 0.25.
Independent sample t-test for equivalence of means: 1,2t 5 2.0, df 5 11,
P 5 0.07; 2,3t 5 0.1, df 5 8, P 5 0.9; 1,3 t 5 2.3, df 5 11, P 5 0.04.
Fig. 5. P levels before and after estimated parturition (7 1 day).
Means were used for individual females with multiple values
within each period.
Am. J. Primatol.
444 / Blanco and Meyer
Our data indicate that successful extraction of
female reproductive hormones can be obtained from
oven-dried fecal pellets collected in the field from
small live-trapped nocturnal lemurs. Lynch et al.
[2003] pointed out that fecal data are ‘‘noisier’’ than
serum or urine data, which is likely owing to
variation in the composition of fecal matter and
bacterial degradation among other factors. On the
other hand, fecal collection obviates the need for
more invasive procedures such as blood withdrawal
that could result in stress-induced variation.
Furthermore, fecal samples contain analytes that
have accumulated between defecation events, thus
potentially providing a more representative ‘‘picture’’ of the reproductive state of an individual. In
contrast, values obtained from other sources such as
serum represent ‘‘snapshots’’ of metabolic and
chemical levels at a given point in time [Bardi
et al., 2003; Whitten et al., 1998].
Our data show a pattern of hormonal variation
that is generally consistent with expectations derived
from published data in other lemur species. Fecal E2
levels increased before or at the day of estrus, and
subsequently decreased, in agreement with results
from urinary E2 measurements in gray mouse
lemurs reported by Perret [2005]. During the final
10 days before parturition, however, fecal E2 levels
were higher than post-estrus values. This finding is
consistent with results from some other lemuroids,
such as captive sifakas (Propithecus verreauxi)
[Brockman et al., 1995], but not wild mongoose
lemurs (Eulemur mongoz) [Curtis et al., 2000].
Elevated levels of E2 shortly before parturition may
stimulate lactogenesis by indirectly enhancing prolactin secretion by the pituitary gland [Hadley,
2000]. However, E2 levels should be interpreted with
caution. Ostner and Heistermann [2003] suggested
that variation in fecal estrogen excretion during the
second half of pregnancy in female red fronted
lemurs (E. fulvus rufus) is related to the sex of the
offspring, with increased estrogen levels occurring
only in females giving birth to males. This may be a
prosimian characteristic, as this phenomenon has
been also reported in ruffed lemurs (Varecia variegata) and tentatively mongoose lemurs as well
[Ostner & Heistermann, 2003].
We also found that fecal P levels started to rise
after estrus in most of the females, although values
were consistently low during ovulation and early
pregnancy (Fig. 2). Similarly, Buesching et al. [1998]
observed that vaginal opening in captive female gray
mouse lemurs occurred when plasma P levels
reached their minimum and considered this hormone to be an unreliable marker of reproductive
state at the time of estrus. Perret [1986] attributed
variation of plasma P levels during the luteal phase
of nonpregnant females to differences in social
Am. J. Primatol.
conditions. We observed higher P values around
the time of parturition (Fig. 3). Those levels started
to decrease after birth, although they remained
elevated above luteal values for at least 2 weeks, a
considerably longer period than that reported for red
fronted lemurs in which low mating concentrations
of fecal progestins were attained within 5 days after
parturition [Ostner & Heistermann, 2003]. It has
been reported that post-partum secretion of P in
females with suckling infants is owing to the
maintenance of the corpus luteum of pregnancy via
increased levels of prolactin associated with lactation
[Knobil & Neill, 2006].
Finally, we observed significant intra- and interindividual variation in both E2 and P throughout the
reproductive season. This may hinder the use of fecal
hormonal analysis as a reliable tool to determine
reproductive condition in the absence of other
supplementary data (e.g. observations of vaginal
morphology, nipple development, abdomen palpation
or behavioral observations). We suggest, however,
that the pattern of variation may be worth studying.
Buesching et al. [1998] and Perret [2005] observed
variability in hormone levels in their studies on gray
mouse lemurs in captivity. Perret [2005] reported
relatively higher and variable urinary E2 levels in
younger individuals at estrus and higher E2 levels
before estrus in female mouse lemurs giving birth to
female-biased litters. Although it is not possible to
test the latter observation under these study conditions, we did find, in agreement with Perret, more
variation in E2 levels around estrus in the younger
females (Table II). Another source of variation in P is
spontaneous resorptions during early pregnancies
that may pass undetected without direct hormonal
sampling. Although Perret [1982] suggested that
early undetected resorptions can shorten interestrous intervals in mouse lemur females, further
hormonal analyses are necessary to corroborate this
proposition. Even perinatal death of the offspring
would trigger the physiological changes associated
with a renewed estrus (e.g. abrupt decreases of
P during apparent pregnancy or right after parturition). In turn, both instances may be correlated
with age, if younger individuals, for example, are
prone to reproductive failures more often than
adults. We report the case of female ‘‘K’’, who
was a young female in 2005 (1-year-old) and was
frequently captured during the reproductive season.
She showed high E2 levels on the day of estrus
(Table I), but experienced an irregular weight
gain profile during pregnancy [personal observation].
Furthermore, she displayed a swollen vagina on
the last day of capture (December 22, 2005, about 5
days after estimated parturition), suggesting that
her offspring, if born alive, had probably died. Her
samples showed relatively low E2 levels before
parturition (Fig. 4) and a sharp decrease in P after
giving birth (values were below the sensitivity of the
Fecal Excreted Steroids in Microcebus rufus / 445
assay). Thus, her hormone ‘‘profile’’ in addition to
body mass data and observations of vaginal morphology provide a more complete picture of her reproductive status. Such findings support the contention
that fecal hormonal analysis can be a useful
supplementary tool to help understand the
pattern of reproductive variation in wild primate
In conclusion, this study represents the first
report of fecal E2 and P levels from wild nocturnal
lemurs as well as the first to use a milling machine to
grind fecal pellets for steroid extraction in a primate.
Fecal E2 and P levels obtained from wild brown
mouse lemur females during the reproductive season
were generally consistent with cytological and
captive data, but intra- and inter-individual variation was found to be significant and widespread
during the reproductive season. The application of
less-invasive techniques to construct endocrine profiles in small-bodied nocturnal lemurs in the wild has
the potential to increase our knowledge of their
reproductive biology by providing additional data
that may not be available through direct observations of wild-trapped individuals. Beyond the analysis of individual hormonal profiles, investigation of
possible sources of variation at the population level,
such as age and litter characteristics (e.g. sex and
size), may enhance our understanding of reproduction and population dynamics of nocturnal lemurs in
the wild.
This project benefited from the help of many
people, particularly Laurie Godfrey and Lynnette
Sievert, who provided advice at every stage. We are
thankful to Anja Deppe for her collaboration with
mouse lemur research as well as Patricia Wright for
support in the field. Additional thanks go to our local
(Centre ValBio-trained) research assistant Victor
Rasendrinirina, the Centre ValBio director Anna
Feistner, Jean Claude Razafimahaimodison, Aimée
Razafiarimalala and other personnel of the Centre
ValBio for logistic support. Fieldwork in Madagascar
was conducted under permission of the Ministry of
Environment, Water and Forests, and the CAFF/
CORE committee and our research protocol was
authorized by the University of Massachusetts
IACUC. Research in Madagascar was facilitated by
staff from the Association Nationale pour la Gestion
des Aires Protégées (ANGAP), the Institute for the
Conservation of Tropical Environments (ICTE,
Stony Brook) and the Madagascar Institute pour la
Conservation des Ecosystèmes Tropicaux, (MICET),
especially its director, Benjamin Andriamihaja. We
thank Jeff Wyatt for veterinary advice. Comments on
an earlier draft of this manuscript by Nancy Forger,
Sylvia Atsalis, Toni Ziegler and Anna Feistner are
also greatly appreciated. This paper was written with
the support of the Rufford Foundation, MMBF/CI
Primate Action Fund and Institute of Biotechnology,
Helsinki to M. B. B. and NSF BCS-0721233 to
Patricia C. Wright, Laurie R. Godfrey and Jukka
Jernvall. Comments from two anonymous reviewers
on previous versions of this manuscript are greatly
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