1
The development of gut microbiota in ostriches
and its association with juvenile growth
Elin Videvall
1
, Se Jin Song
2
, Hanna M. Bensch
1
, Maria Strandh
1
, Anel Engelbrecht
3
, Naomi Serfontein
4
,
Olof Hellgren
1
, Adriaan Olivier
5
, Schalk Cloete
3,6
, Rob Knight
2,7,8
, Charlie K. Cornwallis
1
1
Department of Biology, Lund University, Lund, Sweden
2
Department of Pediatrics, University of California San Diego, La Jolla, CA, USA
3
Directorate Animal Sciences, Western Cape Department of Agriculture, Elsenburg, South Africa
4
Western Cape Agricultural Research Trust, Elsenburg, South Africa
5
Klein Karoo International, Research and Development, Oudtshoorn, South Africa
6
Department of Animal Sciences, Stellenbosch University, Matieland, South Africa
7
Department of Computer Science & Engineering, University of California San Diego, La Jolla, CA, USA
8
Center for Microbiome Innovation, University of California San Diego, La Jolla, CA, USA
Corresponding author
Elin Videvall (elin.videvall@biol.lu.se)
Running title
Development of ostrich gut microbiota
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Abstract
The development of gut microbiota during ontogeny in vertebrates is emerging as an important process
influencing physiology, immune system, health, and adult fitness. However, we have little knowledge of
how the gut microbiome is colonised and develops in non-model organisms, and to what extent microbial
diversity and specific taxa influence changes in fitness-related traits. Here, we used 16S rRNA gene
sequencing to describe the successional development of the faecal microbiota in juvenile ostriches
(Struthio camelus; n = 71) over their first three months of life, during which time a five-fold difference in
weight was observed. We found a gradual increase in microbial diversity with age, an overall
convergence in community composition among individuals, multiple colonisation and extinction events,
and major taxonomic shifts coinciding with the cessation of yolk absorption. In addition, we discovered
significant but complex associations between juvenile growth and microbial diversity, and identified
distinct bacterial groups that had positive (Bacteroidaceae) and negative (Enterobacteriaceae,
Enterococcaceae, Lactobacillaceae) correlations with the growth of individuals at specific ages. These
results have broad implications for our understanding of the development of gut microbiota and its
association with juvenile growth.
Keywords
gut microbiome, struthio camelus, development, succession, colonisation, weight
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Introduction
The gastrointestinal tract of vertebrates is considered to be largely sterile at the time of birth (Perez-
Muñoz et al. 2017; cf. Jiménez et al. 2008) and subsequently colonised by a wide array of micro-
organisms, collectively termed ‘the gut microbiota’. The gut microbial composition during early life has
been shown to have major influences on the health and phenotype of adults through its effects on gut
morphology, metabolism, immune system development, and brain development (Dominguez-Bello et al.
2010; Heijtz et al. 2011; Russell et al. 2012; Cho et al. 2012; Cox et al. 2014). For example, animals
prevented from acquiring gut bacteria suffer from smaller intestines with thinner gut walls, smaller lymph
nodes, a poorly developed immune system, and reduced organ sizes including heart, lungs, and liver
(Gordon & Pesti 1971; Mitsuhiro & Jun-ichi 1994; Macpherson & Harris 2004). Similarly, animals with a
poorly developed gut microbiota have an altered metabolism (Cox et al. 2014) and are more susceptible
to infection by pathogenic bacteria, viruses, and eukaryotes (Sprinz et al. 1961; Inagaki et al. 1996;
Round & Mazmanian 2009). Given the crucial effects of gut bacteria on hosts, it is important to
characterise how, when, and by what microbes the gut is colonised, to examine whether variation in this
process explains differences in host development.
The majority of research on the microbial colonisation of the gut during host development, and its
associated effects on fitness, has been on humans, as well as domesticated and model laboratory animals.
In some animals it has been found that the diversity of the gut microbiota increases with age during
ontogeny, whereas in others the reverse is true. For example, in mice and humans, colonisation is initiated
during birth, where the mother’s vaginal and skin microbiota are important sources of bacteria (Sommer
& Bäckhed 2013; Pantoja-Feliciano et al. 2013; Kundu et al. 2017). Seeding of microbes continues
through lactation, and during the first year of life the human gut microbiome remains relatively simple
with low diversity, and varies markedly across individuals and over time. The gut microbiome
subsequently shifts during weaning towards an adult-like bacterial community and becomes more stable
(Sekirov et al. 2010; Koenig et al. 2011; Yatsunenko et al. 2012). In contrast, in species such as zebrafish
(Danio rerio) and African turquoise killifish (Nothobranchius furzeri), the alpha diversity and richness of
the gut microbiota is highest in neonatal juveniles and subsequently decreases during maturation
(Stephens et al. 2016; Smith et al. 2017). Similar to fish, juvenile birds rely heavily on environmental and
dietary sources for acquiring the initial gut microbes (Lu et al. 2003; Yin et al. 2010). However,
depending on the level of parental care, some bird species may receive significant microbial contributions
from their parents, via for example regurgitation or shared nest environment (Godoy-Vitorino et al. 2010;
van Dongen et al. 2013; Dewar et al. 2017).
Differences in the colonisation of gut microbiota during development have been shown to have
pronounced and highly variable long-term effects on hosts. It has been found that bacterial diversity in the
gut promote host development and growth by enabling greater resource acquisition and preventing
domination by certain bacteria (Ley et al. 2006; Lozupone et al. 2012; Foster et al. 2017). For example,
studies have demonstrated that germ-free animals require a higher calorific intake to attain the same
growth as hosts with a normal microbial diversity (Wostmann et al. 1983; Bäckhed et al. 2004; Shin et al.
2011; Sommer & Bäckhed 2013). Conversely, it has been suggested that a reduced diversity in the gut
microbiota may increase growth and accelerate host development. This idea is supported by numerous
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studies from the agricultural industry where higher growth rates have been achieved in farm animals by
eliminating gut bacteria with antibiotics, a common practice since the 1950s (Gaskins et al. 2002; Dibner
& Richards 2005; Lin et al. 2013). Likewise, supplementing wild animals with antibiotics has also been
associated with positive effects on growth (Potti et al. 2002; Kohl et al. 2017). Pinpointing the exact
mechanisms through which gut microbes influence host growth has, however, been problematic as
antibiotics in some cases increase microbial diversity (Crisol-Martínez et al. 2017; Kohl et al. 2017).
Similarly, some probiotic supplements have led to an increase in animal growth while others are
associated with a reduction in growth (Million et al. 2012; Angelakis et al. 2013). These alternative
predictions and conflicting reports on gut microbiota and animal growth highlight the need for separating
the effects of diversity and specific bacterial groups. For example, different taxa may be associated with
either an increase or decrease in host metabolism and/or intestinal immune responses, which may
engender highly different effects on juvenile development and growth.
In this study, we evaluated the developing gut microbiome of ostrich (Struthio camelus) chicks over time
and in relation to their growth. We performed repeated faecal sampling of individually tagged ostriches in
a research rearing facility from their first week after hatching until 12 weeks of age, which constitutes the
critical developmental phase in this species (Verwoerd et al. 1999; Cloete et al. 2001). Ostriches are the
largest living bird species and, together with other paleognaths, are basal in the phylogeny of birds. They
are a valuable economic resource being farmed for feathers, meat, eggs, and leather, yet have only been
kept in captivity for a short period of time relative to other agricultural animals (Cloete et al. 2012). Their
chicks are highly precocial, allowing them to be raised independently from parents, and they reach sexual
maturity from two years of age. Ostriches also have one of the largest variations in offspring growth rate
across birds, even in controlled environments (Deeming & Ayres 1994; Skadhauge & Dawson 1999;
Bonato et al. 2009; Engelbrecht et al. 2011), and are known to suffer from bacterial gut infections
(Verwoerd 2000; Keokilwe et al. 2015). These traits make the ostrich an excellent organism for
investigating host-microbiota associations, including the effects of gut microbiota on juvenile growth and
development.
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Materials and methods
Experimental setup
Juvenile ostriches were kept under controlled conditions at the Western Cape Department of Agriculture’s
ostrich research facility in Oudtshoorn, South Africa. Chicks were obtained from a batch of artificially
incubated eggs that hatched on Sep 30
th
2014. A total of 234 individuals were monitored from hatching
date until three months of age (12 weeks) in four groups that contained around 58 chicks each at the start
of the experiment. The groups were kept in indoor pens of approximately 4×8 m with access to outdoor
enclosures with soil substrate during the day. To reduce potential environmental variation on the
development of the gut microbiota, all individuals were reared under standardized conditions with ad
libitum access to food and fresh water during the daytime. The chicks received a standardised pelleted
pre-starter ration, and the adult birds were given pelleted breeder ration and were kept in a different area
separate from the chick facility. All procedures were approved by the Departmental Ethics Committee for
Research on Animals (DECRA) of the Western Cape Department of Agriculture, reference no. R13/90.
Sample collection
Faecal samples in this study were collected from chicks during the following ages: week 1, 2, 4, 6, 8, and
12. In addition, we sampled fresh faeces from five adult individuals kept in large outside enclosures. The
sex and age of the adults are not known, but the samples were collected from sexually mature, breeding
individuals. All faecal samples were collected in empty plastic 2 ml micro tubes (Sarstedt, cat no. 72.693)
and stored at -20 °C within two hours of collection. They were subsequently transported on ice to a
laboratory and stored at -20 °C. Detailed sample collection has been described by Videvall et al. (2017a).
Weight measurements of the ostriches were retrieved during each sampling event. At the final time point
(week 12), the smallest ostrich chick weighed 6 kg while the largest weighed 30 kg, representing a five-
fold difference in body mass (mean = 18 kg).
Throughout the course of the trial, a large number of individuals died (n = 72) primarily from suspected
disease, as is common in ostrich rearing facilities (Verwoerd et al. 1999; Cloete et al. 2001). In addition,
10 chicks were randomly selected for euthanization and dissection at 2, 4, 6, 8, 10, and 12 weeks of age (n
= 60), to act as age-matched controls for the diseased individuals. The gut samples of the diseased and
euthanized juveniles have been analysed in a different study (Videvall et al. unpublished data). To
investigate the development and maturation of ostrich microbiomes, the samples used in this study were
those retrieved from euthanized (control) individuals and from individuals that survived the entire period
(n = 71). The faecal samples from the individuals that died from suspected disease were not included.
DNA isolation, library preparation, and amplicon sequencing
We prepared sample slurries for all sample types with guidance from Flores et al. (2012) and
subsequently extracted DNA using the PowerSoil-htp 96 well soil DNA isolation kit (Mo Bio
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