Chimpanzee Diet: Phytolithic Analysis of Feces

Abstract

Most primate populations remain unobservable; therefore, researchers depend on the analyses of indirect evidence encountered at a study-site in order to understand their behavioral ecology. Diet can be determined through the analyses of scats or feeding remains encountered on-site. This allows aspects of their dietary repertoire to be established, which has implications both for conservation efforts (by locating food resources), and for understanding the evolution of hominin diet (if used as referential models). Macroscopic inspection of fecal samples is a common method applied to ascertain a primate population's diet. However, new approaches are required to identify food-items unrecognizable at this level. We applied a dry ash extraction method to fecal samples (N = 50) collected from 10 adult chimpanzees in Kanyawara, Kibale National Park, Uganda and also to plant parts (N = 66) from 34 species known to be included in the diet of this community of apes. We recovered phytoliths in 26 of the 34 plant species. Fifteen phytolith morphotypes were only detected in 14 plant species (termed "distinct" phytoliths). We used these distinct phytoliths to identify plant foods (i.e., that they were associated with) in fecal samples. We then validated findings by checking if the 10 chimpanzees had eaten parts of these plants ∼24 hr prior to fecal sample collection; six plant species associated with five distinct phytoliths had been eaten. Finally, we compared plant foods identified in fecal samples from phytolith analyses with plants that had been identified from macroscopic inspection of the same fecal samples. Findings from phytolith analyses corroborate with those from macroscopic inspection by expanding the total number of plant species identified per fecal sample (i.e., we identified certain plant parts that remained unrecognizable at macroscopic level). This study highlights the potential of phytolith analyses of feces to increase our knowledgebase of the dietary repertoire of primate populations. Am. J. Primatol. © 2014 Wiley Periodicals, Inc.

Full text

RESEARCH ARTICLE
Chimpanzee Diet: Phytolithic Analysis of Feces
CAROLINE PHILLIPS
1
*
,
AND CARLA LANCELOTTI
2,3
1
Department of Archaeology and Anthropology, University of Cambridge, Cambridge CB2 1QH, England
2
CASEs, IMF—Spanish National Research Council (CSIC), Barcelona E‐08001, Spain
3
Department of Communication and Information Technologies, Universitat Pompeu Fabra, Barcelona 08018, Spain
Most primate populations remain unobservable; therefore, researchers depend on the analyses of
indirect evidence encountered at a study‐site in order to understand their behavioral ecology. Diet can be
determined through the analyses of scats or feeding remains encountered on‐site. This allows aspects of
their dietary repertoire to be established, which has implications both for conservation efforts (by
locating food resources), and for understanding the evolution of hominin diet (if used as referential
models). Macroscopic inspection of fecal samples is a common method applied to ascertain a primate
population’s diet. However, new approaches are required to identify food‐items unrecognizable at this
level. We applied a dry ash extraction method to fecal samples (N¼50) collected from 10 adult
chimpanzees in Kanyawara, Kibale National Park, Uganda and also to plant parts (N¼66) from 34
species known to be included in the diet of this community of apes. We recovered phytoliths in 26 of the 34
plant species. Fifteen phytolith morphotypes were only detected in 14 plant species (termed “distinct”
phytoliths). We used these distinct phytoliths to identify plant foods (i.e., that they were associated with)
in fecal samples. We then validated findings by checking if the 10 chimpanzees had eaten parts of these
plants 24 hr prior to fecal sample collection; six plant species associated with five distinct phytoliths
had been eaten. Finally, we compared plant foods identified in fecal samples from phytolith analyses
with plants that had been identified from macroscopic inspection of the same fecal samples. Findings
from phytolith analyses corroborate with those from macroscopic inspection by expanding the total
number of plant species identified per fecal sample (i.e., we identified certain plant parts that remained
unrecognizable at macroscopic level). This study highlights the potential of phytolith analyses of feces to
increase our knowledgebase of the dietary repertoire of primate populations. Am. J. Primatol. 76:757–
773, 2014. © 2014 Wiley Periodicals, Inc.
Key words: ape; fecal; phytolith; plant food‐items
INTRODUCTION
Ascertaining the full range of dietary constitu-
ents of a primate population allows further under-
standing of their feeding ecology. Furthermore, such
work can assist in the identification of habitat types
utilized for food resources within their home range,
which may then assist conservation efforts. More-
over, increased knowledge of the omnivorous diet
of our closest living relatives (Pan troglodytes and
P. paniscus), who range and forage in different
habitats may provide a greater understanding of
hominin diet and its evolution from our last common
ancestor (when used as referential models to deter-
mine both similarities and differences in plant avail-
ability and food‐intake [Copeland, 2009; Hohmann,
2009]). Direct observation of feeding by primates
reveals crucial aspects of their diet. However, for
chimpanzees, such behavioral data are available at
fewer than 25% of current study‐sites, where
populations tolerate human observers at close range
(i.e., they are habituated [McGrew, 2007]).
It is likely that most primate populations will
remain unobservable and there are good reasons for
this. First, a long‐term commitment by researchers
is required to accomplish habituation, due to the
subjects’initial wariness. Indeed, full habituation
may take years to achieve [Bertolani & Boesch,
Contract grant sponsor: Murray Edwards College of Cambridge;
contract grant sponsor: Board of Graduate Studies of University
of Cambridge; contract grant sponsor: Ridgeway‐Venn Travel
Fund; contract grant sponsor: Sir Richard Stapely Education
Trust; contract grant sponsor: Leverhulme Trust
Correspondence to: Evolutionary Studies Institute, University
of Witwatersrand, Wits 2050, South Africa.
E‐mail: caroline.phillips@wits.ac.za
Received 5 August 2013; revised 28 January 2014; revision
accepted 30 January 2014
DOI: 10.1002/ajp.22267
Published online 23 February 2014 in Wiley Online Library
(wileyonlinelibrary.com).
American Journal of Primatology 76:757–773 (2014)
© 2014 Wiley Periodicals, Inc.

2008; Doran‐Sheehy et al., 2007]. Also, some
researchers choose not to habituate chimpanzees
in unprotected areas to maintain their caution
towards human hunters and farmers [Deblauwe &
Janssens, 2008]. Furthermore, there is a rising
concern of the risk of fatal disease transfer from
human observer to subject [Köndgen et al., 2008;
Woodford et al., 2002].
For unhabituated populations, dietary informa-
tion can be obtained from recovered food remains and
macroscopic inspection of fecal samples. The latter is
commonly used [Doran et al., 2002; Julliot &
Sabatier, 1993; Tutin & Fernandez, 1993] and entails
weighing and sluicing samples through fine‐meshed
sieves to remove fecal matrix [McGrew et al., 2009;
Moreno‐Black, 1979]. This process leaves residual
food components visible to the naked eye, which then
can be separated, identified, and counted [McGrew
et al., 1988]. However, food‐items that have been
thoroughly masticated or digested, (e.g., leaves and
“soft‐bodied”insects) may be unrecognizable at this
level of inspection [Phillips & McGrew, 2013] com-
pared to indigestible food‐items (e.g., fruit seeds and
chitinous exoskeleton). Consequently, if researchers
apply this method alone to fecal samples, the full
dietary repertoire of species ingested by a primate
population may remain unknown.
Phytoliths are opal silica microfossils, or
“plant stones”(phyto meaning plant and lith
meaning stone [Baker, 1959]). They are formed
when soluble silica enters the plant during water,
nutrient and other mineral uptake and is then
deposited within cell lumina, in the cells’wall or
in between cells. With evapotranspiration, the
deposited silica solidifies producing silica plant‐cell‐
structure formations, which can be distinctive due to
their size and morphology [Piperno, 2006]. These
formations are used to identify plants, where possi-
ble, to family or genus level [Piperno, 1988; Rovner,
1971].
As a plant decomposes, phytoliths are normally
deposited locally into sediments. Deposited phyto-
liths can dissolve [Cabanes et al., 2011], which has
been shown to maintain equilibrium in concentra-
tions of silica in soils [Farmer et al., 2005], but not all
phytoliths undergo dissolution. As a consequence,
preserved phytoliths that have remained predomi-
nantly undisturbed [Cabanes et al., 2011; Runge &
Fimbel, 2001] have been found in excavated sedi-
ments worldwide. This, along with the ability to
diagnose phytoliths to plant type (e.g., grass), and
potentially to taxon makes phytoliths a powerful tool
to reconstruct paleoenvironments, in particular to
investigate landscape use by human ancestors
[Barboni et al., 2010; Madella et al., 2002; Wang
et al., 2003] and modern humans [Jiang, 1995;
Pearsall, 1994; Piperno, 1994].
By surviving both mastication and digestion,
phytoliths have also been used to study diet.
Phytoliths successfully extracted from dental calcu-
lus [Henry et al., 2010; Lalueza & Pérez‐Pérez, 1994],
including calculus from the extinct primate Gigan-
topithecus blacki [Ciochon et al., 1990], coprolites
[Horrocks & Irwin, 2003; Prasad et al., 2005], and
more recently, from fresh feces [Lancelotti, 2010;
Lancelotti & Madella, 2012] have all provided
valuable insights into dietary intake of various
extinct and extant species. A preliminary study of
phytoliths detected in mantled howler monkey
(Alouatta palliate) feces is provided by Teaford and
Glander [1996], and the potential of analyzing plant
microfossils (including phytoliths) in feces of extant
primates to infer diet is discussed further in Henry
[2012].
The objective of this study was to investigate
phytolith content of feces from 10 adult chimpanzees
of the Kanyawara community in Kibale National
Park, Uganda to assess what can be gleaned about
the diet of a wild ape population using these proxies.
We applied a dry ash extraction method [Lancelotti,
2010; Parr et al., 2001] to fecal samples collected from
the 10 focal individuals. Plant samples collected in
the home range of this ape community were also
analyzed using this method. Firstly, we sought to
establish if phytolith production occurs in the
separate plant parts, and if so, if these parts could
then be differentiated from each other by their
phytolith content and morphology. We then com-
pared phytoliths from fecal samples collected from
the 10 focal individuals to the reference library
created to investigate diet. Finally, we review the
potential of this technique to corroborate with
findings from alternative fecal analysis, in this case
macroscopic inspection. A particular aim was to
detect those parts of plant species seen to be eaten by
the 10 focal individuals (e.g., leaves or piths) that had
not been identified at macroscopic level in the same
fecal samples [Phillips & McGrew, 2013].
METHODS
Study‐Site
Kanyawara is located in the northwestern part of
Kibale National Park (0°130–0°410N, 30°190–30°320E)
in Uganda, at about 1,500 m elevation [Wrangham
et al., 1994]. This mature, mid‐altitude semi‐decid-
uous and evergreen ecotype has swamp, primary, and
regenerating forest that was logged pre‐1992 [O’Dris-
coll Worman & Chapman, 2004]. Dominant trees in
unlogged areas are Parinari excelsa,Celtis gompho-
phylla, and Markhamia lutea [Sekercioglu, 2002],
while the woody shrub Acanthus polystachius dom-
inates previously logged areas [Osborne et al., 2001].
Dominant grasses within the national park include
Pennisetum purpureum,Hyparrhenia spp., and
Cymbopogon nardus [Zanne & Chapman, 2005].
There are distinct wet and dry seasons: May–August
Am. J. Primatol.
758 / Phillips and Lancelotti

and December–February are drier than other months
[Chapman et al., 2001]. We collected data over
162 days between June and December 2008 (exclud-
ing August). Thus, data covered two of the four
months of one dry season (D
1
), all of one wet season
(W) (September–November) and 1 of 4 months of a
second dry season (D
2
). During the study period,
mean monthly rainfall during the dry seasons
combined was 52.7 SE 6.6 mm and in the wet
season was 212.1 SE 65.9 mm. Mean maximum and
minimum monthly temperatures in the dry and wet
seasons were 27.9 SE 0.4°C (range: 18.3–38.0°C;
N¼78 days) and 14.1 SE 0.1°C (range: 0.0–17.0°C);
and 28.3 SE 0.5°C (range: 14.1–43.3°C; N¼84
days) and 14.1 SE 0.1°C (range: 11.9–17.5°C) (C.
Chapman, personal communication).
Data Collection
Focal individuals
The Kanyawara community numbered ca.50
individuals at the time of the study. We followed 10
fully‐habituated adult chimpanzees (five males and
five females) individually from the time that they
arose out of their arboreal bed in the morning until
they made another arboreal bed in the evening (one
focal sample, Martin & Bateson, 2007). Each focal‐
sample (N¼19) lasted up to three consecutive days,
where we followed nine individuals twice (for test and
re‐test purposes). The research adhered to guidelines
as set down by the Division of Biological Anthropolo-
gy, University of Cambridge, and the American
Society of Primatologists’principles for the ethical
treatment of non human primates. The Uganda
Wildlife Authority and Uganda National Council for
Science and Technology permitted data collection on
the Kanyawara chimpanzee community in Kibale
National Park.
Sample Collection
We collected 34 plant species (13 herbaceous
dicotyledons, 18 woody dicotyledons and three
herbaceous monocotyledons) representing 25 fami-
lies to create a reference library. We identified plant
specimens with the aid of permanent staff of the
Kibale Chimpanzee Project. All species have been
observed to be eaten by members of the Kanyawara
chimpanzee community within the last 21 years. We
collected plant parts seen to be eaten as well as parts
not eaten by the Kanyawara community to compare
their phytolith productivity. We saw the 10 focal
individuals eat parts of 30 of the 34 plant species
collected. Leaves, fruit and pith of 12 of these 30 plant
species had not been identified during macroscopic
inspection of fecal samples collected from the same 10
focal individuals [Phillips & McGrew, 2013]. We sun‐
dried all collected plant matter, separated them into
plastic bags, and stored them in airtight containers
with silica gel.
We collected most fecal samples 24 hr after the
first encounter (78% of samples) with each focal
individual. This allowed time for digestion and the
passing of food items we saw the focal individuals eat,
which would then appear in subsequent fecal
samples. We collected all samples in 50 ml centrifuge
tubes and desiccated them with silica gel (to make
sample processing off‐site easier), however, as
phytoliths survive plant decomposition, mastication,
and digestion [Piperno, 2006], degradation of feces is
unlikely to affect phytolithic analysis.
Sample Preparation and Extraction
We separated collected plant parts (N¼66)
accordingly. There were: 34 leaves; 20 fruits (whole);
10 pith or stem sections, and 2 blossoms. We prepared
samples and extracted phytoliths from ashed plant
and fecal samples using Lancelotti’s [2010] dry ash
extraction method (Fig. 1). We did not use hydrogen
peroxide during the extraction process as most
organic matter was burnt away when we ashed
samples at 550°C. Furthermore, this chemical may be
very aggressive on phytoliths where organic matter is
not abundant, which in turn can affect the total
number of identifiable phytoliths [Cabanes
et al., 2011].
We observed extracted phytoliths from both
plant and fecal samples under a transmitted light
microscope at 400. We counted phytoliths along
seven random vertical transects per slide, resulting
in a total of 371 view fields (53 view fields per
transect). This count was based on Van der Veen and
Fieller’s [1982] minimum count of 350 for valid
sample material representation per slide. Using a set
number of view fields per slide provided a general
overview of the relative abundance of phytoliths per
plant part analyzed.
In each sample we only counted phytoliths
judged to be complete (i.e., not broken), meaning
that the shape, margin, texture and where possible,
the location within the plant part could be identified
(e.g., epidermal layer). We counted phytoliths within
silica skeletons (phytoliths of plant cells that are still
articulated) individually, as complete silica skeletons
are less frequently found in sediments due to their
breaking up post‐plant decomposition [Tsartsidou
et al., 2007]. We expected that after mastication and
digestion, fewer complete silica skeletons would be
present in fresh fecal samples compared to those
found in the plant reference samples.
We determined phytolith concentrations using
Albert and Weiner’s [2001] calculation, which pro-
vides a total for the number of phytoliths in 1 g of the
acid insoluble fraction (AIF). This fraction denotes
the remaining dried matter that was not dissolved by
acids during the extraction process. Listing the total
number of phytoliths per gram of AIF (phytoliths/
gAIF) for each plant part allowed intra‐and inter‐
species comparison (Table I).
Am. J. Primatol.
Phytoliths in Chimpanzee Feces / 759

Fig. 1. Protocol for phytolith extraction from plant and fecal samples.
Am. J. Primatol.

TABLE I. List of Plant Species Analyzed and Total Number of Phytoliths Counted Per Slide and Per Gram of Acid
Insoluble Infraction in Each Plant Part (N ¼66)
Species, Plant authority Family Form Group Part Eat
Slide
count Phy/gAIF (10
6
)
Acalypha ornata Hochst. ex A.Rich. Euphorbiaceae THV D Leaf Y 38 7.5
Acanthus polystachius Delile Acanthaceae THV D Leaf N 14 2.8
Aframomum spp. Zingiberaceae THV M Leaf Y 1,584 156.5
Stem Y 0 —
Aneilema aequinoctiale (P.Beauv.) Loudon Commelinaceae THV M Leaf Y 74 7.3
Stem N 0 —
Cordia africana Lam. Boraginaceae TRE D Leaf N 859 169.8
Fruit Y 103 6.8
Stem N 322 63.6
Crassocephalum vitellinum (Benth.) S.Moore Compositae THV D Leaf Y 186 3.3
Blossom N 0 —
Chaetacme aristata Planch. Ulmaceae THV D Leaf Y 1,163 76.6
Fruit N 50 1.6
Dovyalis macrocarpa Bamps Flacourtiaceae SHB D Leaf N 0 —
Fruit Y 0 —
Ficus asperifolia Miq. Moraceae SHB D Leaf Y 2,040 201.6
Fruit Y 24 2.4
Ficus exasperata Vahl Moraceae TRE D Leaf Y 2,251 444.9
Fruit Y 132 4.3
Ficus ottoniifolia subsp.
lucanda (Ficalho) C.C.Berg
Moraceae TRE D Leaf N 2,164 85.5
Fruit Y 178 7.0
Ficus sansibarica subsp. macrosperma
(Warb. ex Mildbr. & Burret) C.C.Berg
Moraceae TRE D Leaf N 229 2.8
Fruit Y 196 38.7
Ficus sur Forssk. Moraceae TRE D Leaf N 288 28.4
Fruit Y 24 <0.5
Hoslundia opposita Vahl Lamiaceae THV D Leaf N 0 —
Fruit Y 0 —
Jasminum spp. Oleaceae CLI D Leaf Y 23 4.6
Stem N 0 —
Lepistemon owariense (P. Beauv.) Hallier f. Convolvulaceae THV D Leaf Y 0 —
Maesa lanceolata Forssk. Primulaceae THV D Leaf Y 232 45.9
Fruit Y 53 3.5
Marantochloa leucantha
(K.Schum.) Milne‐Redh.
Marantaceae THV M Leaf Y 1,868 123.1
Fruit Y 3,466 685.1
Pith Y 5,894 233.0
Mimusops bagshawei S.Moore Sapotaceae TRE D Leaf N 5 <0.5
Fruit Y 0 —
Twig N 0 —
Monodora myristica (Gaertn.) Dunal Annonaceae TRE D Leaf N 461 22.8
Fruit Y 0 –
Twig N 0 —
Neoboutonia macrocalyx Pax Euphorbiaceae TRE D Leaf N 1,264 41.6
Piper capense L.f. Piperaceae THV D Leaf N 244 9.6
Stem Y 0 —
Phytolacca dodecandra L’Hér. Phytolaccaceae THV D Leaf N 0 —
Fruit Y 0 —
Stem N 0 —
Pseudospondias microcarpa (A.Rich.) Engl. Anacardiaceae TRE D Leaf Y 843 166.6
Twig N 0 —
Psychotria mahonii C.H. Wright Rubiaceae TRE D Leaf N 0 —
Fruit Y 0 —
Rubus apetalus Poir Rosaceae THV D Leaf N 0 —
Fruit Y 0 —
Secamone africana (Oliv.) Bullock Apocynaceae CLI D Leaf N 739 48.7
Am. J. Primatol.
Phytoliths in Chimpanzee Feces / 761

We compared findings from phytolith analyses of
20 samples with sub‐samples from the same faecal
sample that had been macroscopically analyzed.
Such a comparison would determine if our micro‐
remains composition corroborated with the macro-
scopic inspection data by increasing the total number
of plant species identified per fecal sample. For cross‐
validation of phytolith findings in fecal samples with
food‐intake of each focal individual, we compared
plant species that we had identified from phytolithic
analysis in fecal samples collected from the focal
individual in question against the plant species that
we had directly seen them eat during their focal
sample. We provide a total percentage of positive
validation‐checks (i.e., plant species in feces had been
eaten by the focal individual at least 22 hr prior to
defecation).
Statistical Analyses
We performed canonical correspondence analy-
sis (CCA; linear covariance matrix) on JMP version
10 to reduce dimensions and provide a smaller
set of derived variables, and to see if phytoliths
detected in particular plant parts could discrimi-
nate plant type as opposed to redundant phytolith
morphotypes we encountered across multiple plant
parts. The first CCA explored if the phytoliths
commonly encountered across multiple plant spe-
cies in our reference library (redundant morpho-
types) could be assigned to a particular plant type:
herbaceous monocotyledon (HM); herbaceous dicot-
yledon (HD); and woody dicotyledon (WD). A second
CCA explored if less common phytolith morpho-
types(foundonlyinselectedplantspeciesana-
lyzed) assigned to a particular plant type; and the
third CCA combined redundant and less common
phytolith morphotypes encountered across the
reference library to ascertain how they all assigned
to the three plant types. We used weighted
averages of phytolith/gAIF of each phytolith type
detected in each plant part analyzed for CCA [Ter
Braak, 1987]. We did not include phytolith mor-
photypes (N¼11) that were only encountered in
parts of a single plant species to minimize zero
loading.
We used a Wilcoxon’s signed‐rank test (one‐
tailed) to compare findings of phytolith totals in silica
skeletons between plant and fecal samples to deter-
mine if mastication and digestion affect the level
of phytoliths articulation (i.e., presence of silica
skeletons). As sample size differed across the three
seasonal periods, we used a Kruskall–Wallis test
(two‐tailed) to determine if change in total number
of phytoliths per gram of AIF extracted from the
fecal samples occurred which may reflect seasonal
change in diet. Statistical analyses were completed
on MINITAB Release 14 where alpha was set at
0.05.
RESULTS
Plant Reference Samples
We observed phytoliths in 26 plant species
(76%), and 39 plant parts (25 leaves; 10 fruit; 2
stems, 1 pith; and 1 blossom) from 19 families. The 10
focal individuals had been observed to eat parts from
19 (73%) of the 26 plant species in which phytoliths
were present. Table I lists these plant species. Ten
species and their parts that had phytoliths which we
saw the 10 focal individuals eat, but which were not
identified during macroscopic inspection of their
scats, are highlighted in bold. The remaining eight
plant species in which we did not recover any
phytoliths were all dicotyledonous (Table I) and
TABLE I. (Continued)
Species, Plant authority Family Form Group Part Eat
Slide
count Phy/gAIF (10
6
)
Fruit Y 0 —
Tabernaemontana pachysiphon Stapf Apocynaceae TRE D Leaf N 0 —
Fruit Y 0 —
Tarenna pavettoides (Harv.) Sim Rubiaceae SHB D Leaf N 0 —
Fruit Y 0 —
Toddalia asiatica (L.) Lam. Rutaceae SHB D Leaf N 600 59.3
Fruit Y 89 17.6
Trichilia splendida A.Chev. Meliaceae TRE D Leaf Y 288 56.9
Triumfetta tomentosa Bojer ex Bouton Malvaceae THV D Leaf Y 113 5.6
Urtica massaica Mildbr. Urticaceae THV D Leaf N 447 88.4
Blos Y 239 47.2
Uvariopsis congensis Robyns & Ghesq. Annonaceae TRE D Leaf N 17,798 502.6
Plant form: terrestrial herbaceous vegetation [THV]; tree [TRE]; shrub [SHB]; and climber [CLB]. Plant group: dicotyledon [D], monocotyledon [M]. Part
eaten by chimpanzee: yes [Y], no [N]. Total number of phytoliths counted per slide, and phytolith concentration—total number of phytoliths found per gram of
acid insoluble fraction (Phy/gAIF) for each plant part analyzed. Plant species (N¼10) and part (N¼11) in bold were not identified during macroscopic
inspection of fecal samples collected from the 10 focal chimpanzees.
Am. J. Primatol.
762 / Phillips and Lancelotti

represented 16 of the 66 plant parts analyzed. We
observed the 10 focal individuals eat parts of all of
these taxa.
We detected phytoliths in the leaves of all three
monocotyledons, and for Marantochloa leucantha,
the fruit had more than double the abundance
of phytoliths (measured in phytolith/gAIF) than
the leaf and pith combined. We saw the 10 focal
individuals eat the leaves of all three species, as well
as the pith and fruit of M. leucantha and Aframomum
spp.
Of the herbaceous dicotyledonous species ana-
lyzed, we detected phytoliths in all leaf parts
(Table I). Abundance of phytoliths was highest for
Chaetacme aristata and Urtica massaica leaves,
although for the latter, the focal individuals ate
only the blossom.
For all woody dicotyledonous species in which
phytoliths were found in multiple plant parts,
abundance of phytoliths was higher in leaves than
other parts, with the exception of Ficus sansibarica
macrosperma, in which the phytolith count/gAIF was
approximately 14 times greater in its fruit than
leaves. Chimpanzees ate leaves as well as fruit of
both F. exasperata and F. asperifolia. Leaves of both
species had a substantially higher abundance of
phytoliths compared to fruits, with F. exasperata
having a phytolith concentration nearly 100 times
higher and F. asperifolia about 84 times higher. No
species had phytoliths only in the fruit parts,
however, this is common finding in other species in
which multiple plant parts have been analyzed
[Piperno, 1988].
Phytolith Morphotypes
We detected 32 phytolith morphotypes in the
26 plant species analyzed; their occurrence across
our reference library is shown in Figure 2. The
nomenclature classifications used for phytolith
identification are from Madella et al. [2005],
Mercader et al. [2009], Piperno [1985, 1988, 2006],
and Runge [1999].
Eight phytolith morphotypes occurred frequently
across plant parts of trees, shrubs, terrestrial herbs
(or “THV”, Malenky et al., 1994) and climbers in our
reference library (SI, Supplementary Material).
These redundant phytoliths were: two epidermal
phytoliths (polyhedral and jigsaw of varied textures);
stellate (star‐shaped) hair‐bases; psilate (smooth)
non‐segmented hairs; and stomatae found in tissue of
upper and lower epidermal layers of leaves and fruit
skins [Mercader et al., 2009; Piperno, 1988, 2006];
mesophyll phytoliths from tissue located below the
epidermis; and tracheids and vascular cells, both
found in the vascular tissue of xylem tracheary
elements [Piperno, 2006]. Redundant morphotypes
made up >60% of the total assemblage of phytoliths
counted in our plant reference library and are shown
in Figure 3a–f.
Redundant phytoliths accounted for 80.9 SE
6.1% of total number of phytolith morphotypes
identified for each herbaceous dicotyledon species.
We detected fewer redundant phytoliths in the three
herbaceous monocotyledon THV species; <30% of
phytoliths in Aframomum spp. and Marantochloa
leucantha parts were redundant. Polyhedral epider-
mal phytoliths, stomatae and non‐segmented, psilate
hairs however, were the only phytolith morphotypes
detected in Aneilema aequinoctiale leaves. Redun-
dant phytoliths accounted for 94.3 SE 3.3% of total
number of phytolith morphotypes identified in each of
the 14 woody dicotyledon species. For all fruit (except
Ficus asperifolia), and leaves of each tree species
(except Ficus spp. and Cordia africana leaf and stem),
polyhedral epidermal phytoliths, mesophyll phyto-
liths, tracheids and vascular cells were the only
phytolith morphotypes we identified.
We found 15 other phytolith morphotypes in
parts of 14 of the 26 plant species (Figure 3g–s, SII,
Supplementary Material). These phytolith morpho-
types are not univocally produced by these species
and can occur in other species of their genus and
across the multiple families represented [Barboni
et al., 2007; Piperno, 1988; Runge & Fimbel, 2001].
However, as one of our aims was to assess whether
the phytolith content of fecal samples reflected the
diet of the 10 focal individuals, we discuss these
morphotypes in the context of our plant reference
collection. They are referred to below as “distinct”
phytoliths for this study (i.e., they are not unique, but
they are distinguishable from other phytolith morpho-
types recovered within the plant reference library).
Forty‐seven per cent of these distinct phytoliths
occurred in plant parts of THV species that we saw
the 10 focal individuals eat. We commonly found a
biconical convex (termed “hourglass‐shaped”below),
segmented hair that had a striated texture (“elongat-
ed elements in a parallel line”Madella et al., 2005) in
the leaves of Maesa lanceolata. We found four types of
globular‐shaped phytoliths of differing textures
(nomenclature of each in SII, Supplementary Mate-
rial) in the leaves of various terrestrial herbs (both
monocotyledon and dicotyledon) and in the leaf of
Cordia africana. We also detected them in the pith
and fruit of M. leucantha. Hat‐shaped phytoliths with
a sinuated (“alternating but uneven concavities and
convexities”Madella et al., 2005) margin and verru-
cate (“irregularly shaped, wart‐like”Madella et al.,
2005) texture were both common in the leaves of
Aframomum spp. and all parts of M. leucantha. Hat‐
shaped phytoliths have been detected in various
families including genera of the Zingiberaceae and
Marantaceae [Chen & Smith, 2013; de Albuquerque
et al., 2013; Piperno, 1985, 1988; Runge, 1999].
Elongate cystoliths (outgrowths of cell walls in
epidermal layers) occurred in the leaf of M. leucantha
and in the leaf and blossom of U.massaica. These
morphotypes have been observed in other species of
Am. J. Primatol.
Phytoliths in Chimpanzee Feces / 763

Fig. 2. Total number of samples and plant parts (leaf, fruit, stem/pith, blossom) in which each phytolith morphotype (N¼32) was observed
(e.g., polyhedral epidermal phytoliths were observed in 20 leaves, 6 fruits, 2 stem/piths and 1 blossom in the 26 plant reference samples
analyzed). Phytolith morphotype, listed by its shape, margin (edges of phytolith), and texture. Nomenclature used from: Madella et al.
[2005]; Mercader et al. [2009] (); Piperno [1988, 2006] (); Runge [1999] ().
Am. J. Primatol.
764 / Phillips and Lancelotti

Fig. 3. Redundant and distinct phytolith morphotypes in plant and fecal samples (Scale 20 mm). Redundant phytoliths: (a) polyhedral
epidermal phytoliths in leaf of Toddalia asiatica;(b) in fruit of Ficus exasperata (1) stomatae (2) stellate hair‐base (3) psilate non‐
segmented hair; (c) vascular cell in the leaf of Crassocephalum vitellinum;(d) mesophyll phytoliths in fruit of Toddalia asiatica;(e) jigsaw
epidermal phytoliths in leaf of Crassocephalum vitellinum;(f) tracheid in leaf of Pseudospondias microcarpa. Distinct phytoliths: (g)
segmented hourglass hair with striate patterning in leaf of Maesa lanceolata;(h) part‐armed, non segmented hair in leaf of Ficus
asperifolia;(i) lanceolate phytolith with a papillate texture in leaf of Acanthus polystachius;(j) non‐segmented (wide) psilate hair in leaf of
Ficus exasperata;(k) hat‐shape, sinuate verrucate sideways view in pith of Marantochloa leucantha;(l) hat‐shape, sinuate psilate
sideways view in leaf of Piper capense;(m) elongate cystolith in blossom of Urtica massaica;(n) globulose in leaf of Marantochloa
leucantha;(o) orbicular globulose/hat‐shaped phytolith, with sinuate margin and verrucate texture in the leaf of Marantochloa leucantha;
(p) in leaf of Aframomum spp. (1) globular, with a psilate texture (2) psilate, non‐segmented hair; (q) bulbous shaped side‐hat type (a)
phytolith in leaf of Ficus sur;(r) opaque perforated platelet in leaf of Acanthus polystachius;(s) elongate cystolith in fruit of Marantochloa
leucantha; Phytoliths in fecal samples not found in plant reference samples: (t) bulbous lanceolate hat; (u) bulbous side hat type (b); (v)
bulbous side hat type (b); (w) armed hair‐base; (x) lacunose, hair‐base; (y) ovate blocky; (z) rectangular, elongate blocky; (aa) cylindroid
with ridge; (ab) armed hair‐base; (ac) elongate tenius lacunose; (ad) elongate, echinate with central ridge; (ae) rectangular, elongate
blocky; (af) clavate granulate; (ag) globular granulate.
Am. J. Primatol.

the Urticaceae family [Piperno, 1988, 2006]. We
detected an orbicular globular/hat‐shaped phytolith
with a sinuated margin and verrucate texture only in
the leaf of M. leucantha. Although we recovered
psilate, non‐segmented hairs in 68% of the 26 plant
species, in F. exasperata fruit they were particularly
wide (25–140 mm). Also, we commonly detected a non‐
segmented hair, which was part‐armed (“having a
granular surface, composed of fine knobs or knots”
Madella et al., 2005) from the base to half‐way up the
hair only in the leaves and fruit of F.asperifolia,. A
bulbous‐shaped side‐hat was associated with only
the leaves F. sansibarica macrosperma, F. sur and
F. exasperata. Similarly, we only observed a single‐
lobate phytolith with base in the leaf of Jasminum
spp. Leaves of two species of plants that we did not
see the focal individuals eat presented distinct
phytoliths: an elongated, lanceolate‐shaped (triangu-
lar) phytolith with papillate (has protuberances
which are rounded or pointed) texture as well as
opaque perforated platelets which had a “Swiss
cheese‐like”appearance in the leaf of Acanthus
polystachius; and psilate, hat‐shaped phytoliths
with sinuated margin in Piper capense leaves. We
also found the latter were in the leaf of M. leucantha.
In the first CCA performed, the first canonical
component (CC1) accounted for 91.1% of variance; the
second (CC2), 8.9% (SIII, Supplementary Material).
For redundant phytoliths, the three plant groups
(herbaceous monocotyledon (HM), herbaceous dicoty-
ledon (HD) and woody dicotyledon (WD)) overlapped,
but tracheid phytoliths separated WD from HD plant
type along CC1 (standardized score coefficients SIV,
Supplementary Material). Plant types were not
clearly separated along CC2; however, coefficients
reveal a positive loading of redundant phytoliths
toward WD. In the second CCA performed using non‐
redundant phytoliths, CC1 accounted for 89.7% of
variance where HM was separated from WD and HD.
Along CC2 which accounted for 10.3% of variation;
saddles and segmented hairs with a psilate texture
were more negatively loaded by HM plant type. With
the inclusion of both redundant and non‐redundant
phytoliths in the third CCA performed, the first CC
accounted for 75.9% of variance; CC2 for 24.1%. The
inclusion of redundant phytoliths did not modify plant
type separation; as with the second CCA, HM was
separated from HD and WD plant types. Saddles and
segmented hairs with a psilate texture were again
negatively loaded by HM plant type. Misclassification
of plant species into plant type was highest for
redundant phytoliths in the first CCA performed
(SIII, Supplementary Material).
Fecal samples
We analyzed fecal samples “blindly”(i.e., diet was
assessed from phytoliths observed in fecal samples
prior to cross‐validating with data on directly
observed food‐intake by the 10 focal individuals). In
order to test if mastication and digestion affect the
silica skeletons present in ingested plant parts
(i.e., mechanical taphonomy—Madella & Lancelotti,
2012), we compared the total number of polyhedral
epidermal phytoliths present per silica skeleton in
plant parts analyzed with those counted in each silica
skeleton present in fecal samples. We chose this
morphotype as it was one of the most frequent in
plant parts analyzed, and was most often observed
in silica skeletons (i.e., were still articulated). No
difference in total number of polyhedral epidermal
phytoliths per silica skeleton in plant samples and in
fecal samples occurred (Wilcoxon’s signed‐rank test:
T¼45.2; P¼0.20, N¼15; median number of polyhe-
dral epidermal phytoliths per silica skeleton ¼9 for
plants, 6 for fecal samples; range: 1–133 in plants; 1–
315 in feces). Therefore, mastication and digestion
did not necessarily break down silica skeletons in
plant parts ingested.
Total number of phytoliths counted/gAIF for fecal
samples ranged from 56,000 to 166 million. No
seasonal change occurred for the abundance of
phytoliths in samples across the late part of the first
dry season (D
1
), wet season (W) and early part of the
second dry season (D
2
) (Kruskal–Wallis H‐test:
H¼0.8, P¼0.67, N
D1
¼10, N
W
¼31, ND¼9). Phyto-
liths were present in all 50 fecal samples with a
sample mean of 8.7 SE 0.6 phytolith morphotypes
(range: 1–17). We identified 39 phytolith morpho-
types, 27 (69%) of which had been identified in the
plant reference samples (Fig. 4, also Fig. 3). The 12
remaining morphotypes are illustrated in Figure 3; of
these, armed hair‐bases and elongate, rectangular
blocky phytoliths were most frequent (in 38% fecal
samples). We detected redundant phytoliths in 94%
of fecal samples with a mean of 4.2 SE 0.3 types per
sample, giving a mean proportion of 55 SE 0.7% of
total number of phytolith morphotypes detected per
sample (range:0–100). Three redundant phytoliths
that had been detected in multiple plant parts
were the most frequently recovered (78% of fecal
samples); non‐segmented psilate hairs; stellate hair‐
bases; and polyhedral epidermal phytoliths.
We excluded 18 samples from further analyses
either because they had no distinct phytoliths or they
had distinct phytoliths, but had been collected on the
first day of each two to three consecutive‐day focal
sample, and so data comparison with directly
observed plant food‐intake was not possible. Of the
32 remaining fecal samples, we detected 13 of the 15
distinct phytoliths from the reference library. Seven
of these distinct phytoliths were rarely encountered
in the fecal samples (each occurred in 5 samples);
only the lanceolate papillate phytolith we recovered
in 14 samples. We did not see the focal individuals in
question eat the parts of plant reference samples in
which we had detected them. The five remaining
distinct phytolith types were frequently recovered in
Am. J. Primatol.
766 / Phillips and Lancelotti

fecal samples, and came from plants which we had
observed the focal individuals eat. Of these the non‐
segmented, part‐armed hair from F. asperifolia was
the most frequently detected (18 samples), where we
saw the focal individuals eat the leaf or fruit up to
42 hr prior to defecation (75% of cross‐validation
checks). We identified the non‐segmented, wide,
psilate hair from F. exasperata fruit in five samples,
which was directly observed to be fed upon up to 29 hr
prior to defecation (60% of cross‐validation checks).
Fig. 4. Total number of phytolith morphotypes (N¼39) found in fecal samples analyzed (N¼50). Phytolith morphotypes categorized into:
redundant (light grey); distinct (charcoal grey); not found in plant reference samples (mid grey); uncommon, but non‐distinct phytoliths
(black).
Am. J. Primatol.
Phytoliths in Chimpanzee Feces / 767

Hat‐shaped phytoliths with a sinuated margin and
verrucate texture we recovered in 12 samples for
M. leucantha pith, which the focal individuals ate
for 43% of cross‐validation checks and up to 51 hr
previously. We found elongate cystoliths in six
samples where Urtica massaica blossom had been
eaten up to 22 hr prior to defection for 33% of cross‐
validation checks. In 16 samples, we found globular
psilate phytoliths in leaves of Aframomum spp.,
Chaetacme aristata, and also in multiple parts of
M. leucantha. All of these plant foods had been eaten
up to 51 hr 44% of cross‐validation checks.
Combining Phytolith and Macroscopic Data
We compared phytolith findings from 20 fecal
samples with species identified during macroscopic
inspection of paired samples. As a conservative
measure, we had seen the focal chimpanzees eat all
plant species identified 51 hr prior to defecation.
Distinct phytoliths identified added up to two more
plant species per fecal sample (sample mean ¼38% of
species identified; SV, Supplementary Material).
These were phytoliths that had been detected in
the leaves, fruit, pith or blossom of five species and
one genus. Of these, blossom of U. massaica, along
with the fruit and leaves of F.asperifolia and leaves of
F. exasperata had been recognized during macroscop-
ic inspection. The leaves of three THV plant foods
(M. leucantha,Aframomum spp., and Maesa lanceo-
lata), pith of M. leucantha, and the fruit from
F. exasperata had not.
DISCUSSION
The eight redundant phytolith morphotypes
represented a high proportion of those identified in
the plant reference samples, in particular in dicoty-
ledons, which is to be expected [Piperno, 2006].
Redundant morphotypes were shared across parts of
several plant taxa. The overlap of plant type in CCA
for these phytoliths appears to reflect their redun-
dancy in diagnostic ability. Plant type separation, in
particular herbaceous monocotyledon from the two
dicotyledon plant types occurred when we subjected
non‐redundant phytoliths to CCA. Few distinct
phytoliths were discriminated by plant type; howev-
er, neutral loading of distinct phytoliths also oc-
curred, as well as large loading weights of non‐
distinct phytolith morphotypes. These factors could
explain why some phytolith types were not assigned
to plant type. Furthermore, small sample size of
species (N¼3) categorized into HM, and multiple
zero values present for non‐redundant phytoliths
may have further constrained our findings from CCA.
Introducing phytolith morphotypes encountered in
other monocotyledon species to our current reference
library for additional CCA, or using an alternative
analytical tool such as cluster analysis or probability
modeling may provide further insights in future for
phytolith and plant type association in the diet of this
ape community.
Of the 15 distinct phytolith morphotypes that we
found in select plant species analyzed, eight of them
occur in parts of Aframomum spp. or M. leucantha.
These monocotyledons were also two of the highest
phytolith‐producers in this study. Studied genera of
their families have shown high phytolith production
[Piperno, 1988, 2006; Runge & Fimbel, 2001], in
particular in their reproductive structures [Piperno,
1989], but this is not a general feature for all
monocotyledons [Hodson et al., 2005]. We found the
globular‐shaped, psilate textured phytolith in the
leaves of various species, but also in the pith and fruit
of M. leucantha plus the pith of Aframomum spp.
Chen and Smith [2013] noted that globular phytoliths
were small in species of Zingiberaceae, as was
observed in this study for Aframomum spp. when
compared to M. leucantha plant parts. This phytolith
was abundant in samples from East‐African tropical
forests [Barboni et al., 2007], and along with the other
described globular phytoliths, occurs in various
tropical dicotyledon leaves and seeds, but in fewer
tropical herbaceous monocots [Piperno, 1988]; how-
ever, they occur in the reproductive structure of most
families of Zingiberales [Chen & Smith, 2013].
Finding hat‐shaped phytoliths in M. leucantha
was expected as they have been detected in other
species of the Marantaceae family [Chen & Smith,
2013; Piperno, 1989], but they are also noted in other
plant families [Kealhofer & Piperno, 1998]. Identify-
ing their texture and size may further assist in
taxonomic diagnosis. Hat‐shaped phytoliths with a
verrucate texture have been detected in external
fibers of vascular bundle sheaths in species of
Maranta (de Albuquerque et al., 2013). We found
hat‐shapes with a verrucate texture were of similar
size in M. leucantha and Aframomum spp., but those
with a psilate texture were smaller in P. capense vs.
M. leucantha (6 mm vs. 11 mm).
Having compared phytolith morphotypes in the
leaves and fruit of five Ficus species, hair‐bases and
psilate hairs occurred in both parts for most. These
redundant phytoliths have been found in other fig
plants [Piperno, 1985; Tsartsidou et al., 2007], but
not the bulbous side‐hat type (a) detected in the leaf of
three of the five Ficus species; it may be that it is
specific to regions of Africa. The particularly wide
psilate hair found only in the fruit of Ficus exasperata
is also not recorded for other fig species. Nodular‐
textured phytoliths, which have a similar appearance
to the non‐segmented armed hairs we found in F.
asperifolia fruit and leaf, have been recorded for F.
opposita in Australia [Wallis, 2003]. Armed hairs
have also been recovered in Ficus species of South-
east Asia [Kealhofer & Piperno, 1998]. Although only
recovered in one of the Ficus species analyzed, armed
hairs occur in other genera of the Moraceae family
[Kealhofer & Piperno, 1998].
Am. J. Primatol.
768 / Phillips and Lancelotti

Phytoliths studied in parts of Aneilema spp. have
revealed platelets and psilate flake phytoliths in
West African species [Eichhorn et al., 2010]. It is yet
to be determined if the hourglass‐shaped hair with a
striated texture found in Aneilema aequinoctiale in
this study are found in other species of this genus. In
general, species of Commelinaceae are high phytolith
producers, in which “taxonomically significant mor-
photypes”[Eichhorn et al., 2010] can occur.
Phytoliths are not observed or are uncommon in
the families of the plant species analyzed in this
study in which phytoliths were not detected [Piperno,
1988, 2006]. However, silica concentration, which is
used as an indicator of phytolith productivity, can be
wide‐ranging for members of these families [Hodson
et al., 2005], therefore, other genera in these families
may be high producers, and the low phytolith
productivity in species analyzed may not necessarily
reflect a general pattern.
Distinct Phytoliths Versus Cross‐Validation
Checks
Certain phytoliths associated with plant parts in
the reference library that we did not see the focal
individuals eat during the focal sample in question
were detected in a small number of fecal samples.
There may be a number of reasons for this: First,
some of the plant species we observed them eat may
have been misidentified. Second, certain phytolith
morphotypes may have been misidentified due to: a
“weathering”effect caused by partial dissolution
during the extraction process [Cabanes et al.,
2011]; the fact that we were unable to rotate
phytoliths on the slide (Fig. 1); or some morphotypes
having a similarity to other phytolith morphotypes.
Third, the focal individuals ate the plant part in
question when they were out of view, which was
sample mean ¼4.06 SE 0:36 hr (N¼19 focal sam-
ples) prior to collection of the 50 fecal samples.
Finally, these phytoliths occurred in other plant
species or parts that were not included in the plant
reference library analyzed.
We saw the 10 focal individuals eat parts from
eight non‐analyzed plant species during focal sam-
ples. However, they account for only 23% of cross‐
validation checks of food‐items eaten during each
focal sample period (N¼19 focal samples). Misiden-
tification is possible, but it is highly probable that
unobserved feeding bouts occurred in the time the
focal individuals were out of view. These gaps could
account for the presence of these distinct phytoliths.
All three distinct phytoliths found in either the leaves
of Piper capense or Acanthus polystachius could:
(a) reflect non‐observed consumption of their leaves,
(b) signal the inclusion of pith for A. polystachius
observed to be eaten in four feeding bouts by four of
the chimpanzees 13 to 43 hr previously; or (c) come
from other non‐analyzed plant species known to be
included in the diet of the Kanyawara chimpanzee
community. Further analyses of all of these plant
species and their parts should confirm this.
Direct geophagy (seeking and eating soil frag-
ments) or indirect geophagy (soil particles attached to
plant parts eaten) by focal individuals can account for
any recovered phytoliths from plant species that are
not included in our reference library. We witnessed
seven bouts of geophagy (soil consumption) by four
males and four females 13 to 51 hr prior to fecal
sample collection. This may have contributed to the
phytoliths found in fecal samples, having potentially
been present in soil fragments ingested. We cross‐
validated observed bouts of geophagy with phytoliths
detected in fecal samples. Of the 12 phytolith
morphotypes that were not identified in the plant
reference samples, only the lacunose hair‐base was
present in a fecal sample that had been deposited
after geophagy. We also detected three other mor-
photypes that were not identified in the plant
reference samples. These were: (1) rectangular
elongate blocky; (2) armed hair‐base; and (3) bulbous
lanceolate hat. Each of these occurred in fecal
samples (N¼13) collected either before geophagy,
or during focal samples in which we did not see
geophagy. Therefore, geophagy could not account
solely for the sourcing of 92% of the additional non‐
plant reference sample phytolith morphotypes
detected.
In terms of frequency, blocky phytoliths were one
of the highest occurring phytolith morphotypes in
fecal samples. We found none in plant parts from our
reference library. They have been found in the stem
tissue and leaf tissue of various miombo woodland
plants in Mozambique, which include members of
families in which our select plants analyzed also
belong to (e.g., Annonaceae, Euphorbiaceae, Rubia-
ceae in the stem tissue and Flacourtiaceae in the leaf
tissue) [Mercader et al., 2009] and are associated with
shrubs and forbs [Blinnikov et al., 2001]. This may
indicate that they occur in plants or parts that were
eaten but not yet analyzed.
Of the five distinct phytolith morphotypes that
we found in fecal samples, and which occur in plant
species that we saw the focal individuals eat, two are
associated with leaves, pith or fruit of species in both
Marantaceae and Zingiberaceae; one with another
terrestrial herb; and two were hair phytoliths
recovered in the leaves of two different fig species.
This result is encouraging as the identification of
foods such as pith and leaf of THV that are not easily
detected during macroscopic inspection of feces
[Phillips & McGrew, 2013]. Such foods may therefore
be indicated with the study of phytoliths. Using
phytolith morphotypes to differentiate between Ficus
species eaten and potentially their parts also high-
lights the value of analyzing this microfossil, as this
can be difficult to achieve during macroscopic
inspection.
Am. J. Primatol.
Phytoliths in Chimpanzee Feces / 769

A possible contamination issue affected two plant
samples (Aframomum spp. and Urtica massaica leaf
parts) in which cross‐shaped and saddle phytoliths
were recovered. As these morphotypes are normally
found in grasses this discrepancy may indicate that
extraneous matter containing grass‐associated phy-
toliths was still present on these two samples [Albert
et al., 2007], or that the phytoliths detected had a
similar morphology, but were non‐grass phytoliths
and were misidentified. We also found saddle
phytoliths in 10 fecal samples (range: 1–5 per sample)
which may be the result of geophagy, or the
consumption of grass by the focal individuals. Within
Kibale National Park, shrubs and THV included in
the diet of the Kanyawara chimpanzees have been
recorded at grassland sites, and in open areas which
also contain grass species (usually disturbed sites
resulting from human activity) [Duncan & Duncan,
2000; Zanne & Chapman, 2005]. Dominant grasses
in Kibale National Park are from the Panicoideae
sub‐family [Lang et al., 1962]. Phytolith work on
Panicoideae grass species indicate cross‐shaped
phytoliths occur in high proportions, but saddles
more so in species of the Chloridoideae sub‐family
[Bremond et al., 2007; Twiss et al., 1969]. Grass plant
parts may have been harvested and simultaneously
eaten by a chimpanzee feeding on other terrestrial
plants. Parts of Pennisetum purpureum and Zea mays
have been observed to be eaten by members of the
Kanyawara chimpanzee community. We did not see
focal individuals eat grass, except one adult female
who ate the pith of Zea mays during a crop‐raiding
bout. This grass‐intake was on the third day of
the focal sample, and so subsequent fecal samples
were not collected for analysis. No grass parts were
visible to the naked eye in any of the fecal samples
analyzed.
Applying the dry ash extraction method to plant
samples revealed phytoliths in 10 of the 12 plant
species that were part of the observed diet, but that
had not been identified at macroscopic level. Nine of
the 10 distinct phytoliths were associated with leaves
or pith from seven terrestrial herbs (SI, Supplemen-
tary Material). The pith or leaf of four of these plant
species had been seen to be eaten by the focal
chimpanzee in question within the previous 51 hr of
fecal sample collection, and made up 3% of the total
feeding observed for the 10 adult chimpanzees.
Having been observed in fecal samples, distinct
phytoliths added up to two species to the list of those
identified per fecal sample during macroscopic
inspection. Phytolith analysis therefore corroborated
macroscopic inspection. The financial cost of applying
the dry ash extraction method to all plant and fecal
samples analyzed was also relatively low compared to
wet oxidation. This study illustrates that a multiple‐
method approach to fecal analyses need not be
expensive in order to establish plant foods included
in the diet of an ape population.
If most primate populations remain unhabitu-
ated, and also unprotected from human hunters, they
are potentially exposed to fatal zoonotic diseases,
and can experience habitat encroachment by local
human populations. For this reason, the high level of
precision in understanding their dietary habits
offered by such a multiple‐method approach could
assist conservation efforts for these populations.
Analyses of phytoliths in feces can be used to
illustrate and to monitor, for example, the impact
of habitat encroachment by humans on primate
populations. As the loss of habitat reduces wild‐
food plant availability to a population, they may
incorporate non‐wild food‐items into their dietary
repertoire [Campbell‐Smith et al., 2010; Hockings
et al., 2009]. Phytoliths have been identified from
various crops such as Musa spp., Oryza spp.,
Saccharum spp., Triticum spp., Hordeum spp., and
Zea mays [Ball et al., 1999; Harvey & Fuller, 2005;
Piperno, 1984, 2006] and have been shown to resist
digestion processes [Baker et al., 1961; Brochier
et al., 1992; Shahack‐Gross, 2011]. Therefore, their
associated phytolith morphotypes may be detected in
feces.
Furthermore, habitat use can be reconstructed
from phytoliths identified in feces in order to assist
in the identification of important food resources.
Distinct phytoliths associated with Aframomum spp.,
Acanthus polystachius,Marantochloa leucantha,
Maesa lanceolata, Piper capense and Urtica massaica
detected in feces in this study indicate the 10 focal
chimpanzees utilized patches of terrestrial herba-
ceous vegetation (patch size unknown) from areas
with dicotyledonous trees. By reconstructing the
habitats used by extant chimpanzee populations
we can infer information on plant food availability
and diet, thus providing a valuable perspective
(i.e., highlight similarities and differences) to then
interpret potential plant foods that would have
been available to hominin ancestors [Copeland,
2009].
Following on from this preliminary study, efforts
should focus on the expansion of the phytolith
reference library for Kanyawara flora, in particular,
parts of all species known to be included in the diet of
this chimpanzee community, but also plants that are
dominant in the various habitat types found within
their home‐range (including grasses). This extension
should increase current understanding of phytolith
morphotype‐plant association for Kanyawara, and
also highlight any potential misidentification of
phytolith morphotypes. Furthermore, the expansion
of our reference library may reveal which plant taxon
the 12 phytolith morphotypes detected only in fecal
samples come from. Particular attention should be
paid to plant species in which either the leaf, pith or
stem are eaten. These are predicted to appear either
pulverised or fully‐digested in feces at macroscopic
level.
Am. J. Primatol.
770 / Phillips and Lancelotti

Conclusion
Phytolith research corroborates with findings of
primate feces using macroscopic inspection and this
preliminary study advocates for the use of phytoliths
in feces as a diagnostic tool to help determine the
dietary repertoire of primates. Such analyses can
therefore, be done to investigate diet of unhabituated
populations. The building of a plant reference library
is possible through collection of plant foods that have
been observed to be eaten; have been identified from
macroscopic inspection of scats or the study of feeding‐
remains encountered on‐site; or from data available on
plant foods eaten by another population of the study
species. Adding non‐food plants, such as dominant
species present in different habitat types is advanta-
geous as they provide perspective into which redun-
dant and distinct phytolith morphotypes occur across
both foods and non‐foods within the reference library;
their addition can also clarify phytolith morphotypes
not included in the diet of the study population.
Analyses of soil samples may help to identify
phytoliths recovered in feces that are not found in
plants in the reference library (as a result of direct or
indirect geophagy), and give information on phytolith
productivity in soils across the home range. By
applying this bioarchaeological method, and using
extant primates as proxy for human ancestor dietary
practices, this line of research also falls under primate
archaeology [Haslam et al., 2009]. Of course, identifi-
cation of phytoliths recovered in any sample material
may not necessarily lead to identification of plants to
specific plant taxa, but still, as shown from this study,
they can provide insight into habitat utilized for food
resources by extant primates and expand dietary
knowledge from feces of a study population that is not
attainable from other methods of analysis.
ACKNOWLEDGENTS
The authors thank: Uganda Wildlife Authority,
Uganda Council for Science and Technology, and
Makerere University Biological Field Station for
permission to live and work in Kibale National Park;
Richard Wrangham and Martin Muller of the Kibale
Chimpanzee Project for permission to work under the
project and generous logistical support; all staff of the
Kibale Chimpanzee Project for help in data collection
and for identification of plant samples; Louise
Butterworth, Catherine Kneale, Tamsin O’Connell,
Chris Rolfe and Steve Boreham, University of Cam-
bridge for assistance during laboratory analysis; and
William McGrew for guidance and helpful comments
during the research. We thank Tea Jashashvili, Mike
Huffman and four anonymous reviewers for helpful
comments on the manuscript.
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