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\begin{document}

\title{Microbial DNA extraction method for avian feces and preen oil from
diverse species}



\author[1]{Austin Russell}%
\author[1]{Margaret  Kenna}%
\author[2]{Alex Huynh}%
\author[1]{Amber Rice}%
\affil[1]{Lehigh University}%
\affil[2]{Desales University}%


\vspace{-1em}



  
  \date{May 16, 2024}


\begingroup
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\selectlanguage{english}
\begin{abstract}
As DNA sequencing technology continues to rapidly improve, studies
investigating the microbial communities of host organisms (i.e.,
microbiomes) are becoming not only more popular but also more
financially accessible. Across many taxa, microbiomes can have important
impacts on organismal health and fitness. To evaluate the microbial
community composition of a particular microbiome, microbial DNA must be
successfully extracted. Fecal samples are often easy to collect and are
a good source of gut microbial DNA. Additionally, interest in the avian
preen gland microbiome is rapidly growing, due to the importance of
preen oil for many aspects of avian life. Microbial DNA extractions from
avian fecal and preen oil samples present multiple challenges, however.
Here, we describe a modified PrepMan Ultra Sample Preparation Reagent
microbial DNA extraction method that is less expensive than other
commonly used methodologies and is highly effective for both fecal and
preen oil samples collected from a broad range of avian species. We
expect our method will facilitate microbial DNA extractions from
multiple avian microbiome reservoirs, which have previously proved
difficult and expensive. Our method therefore increases the feasibility
of future studies of avian host microbiomes.%
\end{abstract}%



\sloppy


Manuscript Category: Research Article

Title: Microbial DNA extraction method for avian feces and preen oil
from diverse species

Running Title: Avian Microbial DNA Extraction Method

\emph{Authors} : Austin C. Russell\textsuperscript{1,a}, Margaret A.
Kenna\textsuperscript{1}, Alex Van Huynh\textsuperscript{2}, and Amber
M. Rice\textsuperscript{1}

\emph{1. Department of Biological Sciences, Lehigh University,
Bethlehem, Pennsylvania, 18015, USA}

\emph{2. Department of Biology, DeSales University, Center Valley,
Pennsylvania, 18034, USA}

\textsuperscript{a}Corresponding author; email: aar420@lehigh.edu

All of the authors on this paper have no conflicts of interest.

\emph{}

\emph{\textbf{Abstract}}

As DNA sequencing technology continues to rapidly improve, studies
investigating the microbial communities of host organisms (i.e.,
microbiomes) are becoming not only more popular but also more
financially accessible. Across many taxa, microbiomes can have important
impacts on organismal health and fitness. To evaluate the microbial
community composition of a particular microbiome, microbial DNA must be
successfully extracted. Fecal samples are often easy to collect and are
a good source of gut microbial DNA. Additionally, interest in the avian
preen gland microbiome is rapidly growing, due to the importance of
preen oil for many aspects of avian life. Microbial DNA extractions from
avian fecal and preen oil samples present multiple challenges, however.
Here, we describe a modified PrepMan Ultra Sample Preparation Reagent
microbial DNA extraction method that is less expensive than other
commonly used methodologies and is highly effective for both fecal and
preen oil samples collected from a broad range of avian species. We
expect our method will facilitate microbial DNA extractions from
multiple avian microbiome reservoirs, which have previously proved
difficult and expensive. Our method therefore increases the feasibility
of future studies of avian host microbiomes.

\emph{Keywords: microbiome, Aves, uropygial gland, bacterial symbionts,
feces}

\emph{}

\emph{Introduction}

Animal microbiomes have received increased attention in recent years.
Further, the natural microbial communities that live on a host organism
are known to positively impact health, pathogen resistance, digestive
capabilities, and aid in other natural bodily functions. Environment,
dietary niche, and the health of a host all influence the diversity and
abundance of different bacterial species present in a host organism.
Avian species are globally widespread and play many important roles in
their ecosystems. Understanding the factors that contribute to health
and fitness in birds has become especially important in recent decades
due to the rapid decline in bird populations worldwide. Thus, an
understanding of the different microbiomes present within and across
avian species provides valuable insights about avian health, ecology,
evolution, and conservation. A necessary component of characterizing
such microbiomes is the accurate and effective identification of
microorganisms from specific reservoirs.

Although host species possess many reservoirs for microbial species, one
of the most important is the gut. Avian gut microbiomes and their
relationship to the behavior and fitness of the host have been of great
interest over the past two decades. Specifically, the microbiota that
exist in the gut of a bird have been linked to behavior, cognition,
metabolic function, and digestion. To study the gut microbiome, most
scientists extract bacterial DNA from fecal samples. Although fecal
samples do not perfectly represent the gut microbiome without bias, most
microbiomes have been characterized using bacterial DNA extracted from
fecal samples because it allows the characterization of the bacteria
present in the stomach and along the digestive tract as well.

In addition to gut microbiota, preen gland microbiota are important for
birds . The avian preen gland (or uropygial gland) is located at the
base of the tail and produces oils that help birds clean themselves,
protect feather health, and potentially aid in fighting against
pathogenic bacteria. Birds stimulate this gland with their beaks, and
then spread the resulting oil throughout their feathers in a behavior
called preening. Besides feather health, chemicals from preen oils vary
between species and can mediate communication between individuals in
many contexts, including species recognition and mate choice.
Additionally, preen oil chemistry has been shown to vary with seasons,
aggression levels, and reproductive success. Interestingly, recent
studies have shown that preen oil contains bacteria , which play a role
in the synthesis of the chemical compounds found in the oil. Thus, both
gut and uropygial gland microbiota play important roles in avian life.

To characterize the microbiomes present within a species, microbial DNA
must be extracted from samples collected from different reservoirs. To
facilitate comparisons of the microbiome communities present in
different reservoirs, the field would benefit from a microbial DNA
extraction method that is effective for different sample types. However,
within avian hosts, multiple sample types present particular
difficulties. Specifically, microbial DNA extractions from both avian
fecal material and preen oil present several challenges. Although feces
contains abundant bacteria, avian and reptilian organisms combine their
feces with urine, which contains minimal bacteria and therefore lowers
the overall concentrations of the bacteria present in fecal samples.
Further, urine contains PCR inhibitors, such as uric acid, which can
impede bacterial DNA detection. Additionally, preen oil displays low
bacterial abundance\textsuperscript{42} and exhibits antimicrobial
properties, leading to an expectation of even lower bacterial yields
from preen gland versus avian fecal samples\textsuperscript{61}. The
preen oil microbiome has only recently begun to receive attention in the
literature and there are no standardized methodologies for preen gland
microbial DNA extraction.

Most studies on microbiomes have used commercial DNA extraction kits
that are used on other sources of microbial DNA such as human feces or
soil. For avian fecal matter however, DNA extractions from commercial
kits often do not produce visible bands on agarose gels complicating
verification of extraction success. For example, Eriksson et al. (2017)
compared the performance of six different commercial DNA extraction kits
using mallard duck (\emph{Anas platyrhynchos} ) feces and few bands were
visible on gel images. Additionally, the commercially available fecal
extraction kits can be quite expensive (Table 1).

Thus, there is a need for more effective, efficient, and inexpensive
methodologies for extracting microbial DNA from multiple reservoirs.
Here, we present a new method that effectively and consistently extracts
bacterial DNA from both fecal and preen oil samples collected across a
wide range of avian species.

\emph{Materials and Methods}

\textbf{1. Fecal and Preen Oil Sampling}

We obtained fecal samples from a total of 25 individual birds across 15
species (14 genera, 12 families, and 7 orders (Table 2). To capture all
but one of the Passerines used in this study, (excluding the American
crow (see Table 2)), we used mist nets at feeders in Northampton and
Lehigh Counties in Pennsylvania, USA (U.S.G.S. Federal Banding Permits
23810 to AMR and 24256 to AVH; Pennsylvania Banding Permits 103 to AMR
and 49864 to AVH). Fecal collections require careful handling to ensure
the preservation of DNA. We briefly held each bird individually in a
cage containing a clean cage liner until they defecated, which usually
took no longer than several minutes. We then immediately collected the
feces using tweezers. We also collected fecal samples from two
budgerigars (\emph{Melopsittacus undulatus} ) housed in a local pet
store (PetSmart in Bethlehem, Pennsylvania, USA). Tweezers were cleaned
with ethanol before and after each use. We transferred the feces into a
1.5 mL microcentrifuge tube containing 100\% ethanol and stored the
samples in a -80 \selectlanguage{ngerman}°C freezer until microbial DNA extraction. Although
ethanol kills the living bacteria, it preserves all the DNA present in
the sample so that any bacteria present can be detected even after
freezer storage\textsuperscript{68}.

Additionally, in cooperation with the Wildlands Conservancy Nature
Preserve in Lehigh County, Pennsylvania, we obtained fecal samples from
captive birds including the American crow (\emph{Corvus brachyrhynchos}
), plus several species spanning four additional orders (Table 2):
red-tailed hawk (\emph{Buteo jamaicensis} ), eastern screech owl
(\emph{Megascops asio} ), turkey vulture (\emph{Cathartes aura} ), and
rock pigeon (\emph{Columba livia} ). The Wildlands Conservancy samples
were collected as the individuals were seen defecating in their
enclosures. The collections were placed in fresh Ziploc® bags rather
than in ethanol-filled tubes and placed in a freezer prior to
extraction. To collect from the Canada goose (\emph{Branta canadensis}
), we closely observed wild geese on the DeSales University campus. When
we saw defecation, we used sterile tweezers to lift a small portion of
the fresh fecal samples off the ground and placed them into individual
tubes of ethanol. Because the Canada goose produces larger feces than
the other species we sampled, we were careful to avoid any white
portions of these fecal samples, with the goal of reducing the amount of
uric acid in the samples.

We collected preen oil from a total of 12 individual birds, from 12
passerine species, spanning 10 genera, 8 families and 1 order (Table 3).
All these birds were captured in mist nets at bird feeders in Lehigh
County, Pennsylvania, USA (U.S.G.S Federal Banding Permit 24256 and PA
Game Commission Banding Permit 49864). We cleaned the uropygial gland of
each bird and temporarily cleared away any nearby feathers by gently
applying 100\% ethanol with a cotton ball . We then used small, sterile
forceps to gently squeeze the gland. Once preen oil was secreted from
the uropygial gland, we used a capillary tube to collect a small oil
sample (\textasciitilde{}1-2 mg). After collection, the bottom of the
capillary tube was placed in a 1.5 mL microcentrifuge tube and stored in
a -80 °C freezer prior to microbial DNA extraction.

All capture and sampling procedures were approved either by Lehigh
University's Institutional Animal Care and Use Committee (Protocol
\#237) or by DeSales University's Institutional Animal Care and Use
Committee (Protocol \#1).

\textbf{2. DNA Extractions}

To consistently extract microbial DNA from our avian fecal and preen oil
samples, we revised the original Applied Biosystems\selectlanguage{english} PrepMan\selectlanguage{english} Ultra
Sample Preparation Reagent method for extractions from bacterial
samples, adding a bead beating step. For our fecal samples, we first
removed approximately 0.15 g feces from each sample, taking care to
avoid collecting any of the white uric acid. After pouring any ethanol
off the fecal sample, we placed the sample in a weigh boat in a fume
hood for about one minute to allow the evaporation of excess ethanol. We
then transferred the fecal sample into a 2 mL Fisherbrand\selectlanguage{english}~Free-Standing
Microcentrifuge Tube with Screw Cap that had been filled with
approximately 100 \selectlanguage{greek}µ\selectlanguage{english}L of 460 nm acid-washed glass beads (Sigma-Aldrich,
Inc., St. Louis, Missouri, USA) and 200 \selectlanguage{greek}µ\selectlanguage{english}L of PrepMan Ultra Sample
Preparation Reagent (Thermo Fisher Scientific, Waltham, Massachusetts,
USA). For preen oil samples, we placed the entire capillary tube tip
(containing the preen oil sample) into a 2 mL microcentrifuge tube
containing the same volumes of acid-washed glass beads and PrepMan
Ultra Sample Preparation Reagent as used with the fecal samples. Both
types of sample mixtures were homogenized using a Mini-Beadbeater
(BioSpec Products, Bartlesville, Oklahoma, USA) for 1 minute on high,
and then placed in a water bath for 10 minutes at 100\selectlanguage{ngerman}°C. For preen oil
samples, we removed the capillary tube tip after the water bath step, at
which point the oil sample was no longer observable in the tip of the
capillary tube. For both sample types, we then centrifuged samples at
14,000 rpm for 2 minutes and pipetted off the supernatant. This
supernatant was then used as the DNA extract for PCR. The volume of the
final DNA extract samples was \textasciitilde{}75\selectlanguage{greek}µ\selectlanguage{english}L, depending on how
much could be pipetted off without collecting the acid-washed beads. To
create a negative control, we followed this protocol using 100 uL of
water instead of a fecal sample. For a positive control, a colony
of\emph{Escherichia coli} was prepared and used for extraction and PCR
amplification.

3. \textbf{PCR Amplification of microbial 16S rRNA}

To assess the success of our microbial DNA extraction method, we
performed PCR amplification of hypervariable regions of the 16S rRNA
gene (V3-V4 regions) using our DNA extracts from both fecal and preen
oil samples. The 16S rRNA gene is commonly used to identify bacterial
taxa and quantify microbial diversity. This gene is present in all
bacteria and its highly conserved nature coupled with species-specific
regions of variation allows for identification of different clades of
bacteria.

For most of our DNA extractions from fecal samples, we first mixed 1 \selectlanguage{greek}µ\selectlanguage{english}L
of the concentrated fecal DNA sample with 99 \selectlanguage{greek}µ\selectlanguage{english}L of nuclease free water.
If the fecal sample used for microbial extraction was less than 0.1 g,
we used a non-diluted DNA extract sample. Due to the small volume of
collected preen oil samples, \textasciitilde{}1-2 mg, we did not dilute
the preen oil DNA extracts.

We used a total PCR volume of 20 uL, containing master mix, GC enhancer,
forward and reverse 16S rRNA primers, DNA, and water. Specifically, each
reaction included 10 uL Platinum II Hot-Start Green PCR Master Mix (2X)
from Invitrogen (Waltham, Massachusetts), 4uL of the Platinum GC
Enhancer included with this master mix, 2uL of nuclease free
H\textsubscript{2}0, and 2uL of our DNA extraction. To amplify the V3-V4
region of the 16S rRNA gene, we also included 1uL each of 25uM 341F
(CCTACGGGNGGCWGCAG) and 806R (GACTCHVGGGTATCT-AATCC) 16S rRNA primers.

Optimized PCR conditions included an initial denaturation step at 95\selectlanguage{ngerman}°C
for 2 min, followed by 30 cycles of denaturation at 95°C for 30 seconds,
annealing at 55°C for 30 seconds, extension at 68°C for 1 min, and a
final elongation at 68°C for 2 min. To confirm successful PCR
amplification, we ran each sample on a 2\% agarose gel at 140 V for 50
minutes and verified the presence of bands visually. The expected
product size was \textasciitilde{}430 bp. For any extractions that did
not show a band on the first agarose gel, we either ran the same PCR
product on an additional agarose gel, or we ran a new PCR reaction using
the original DNA extract. We therefore did not extract DNA multiple
times from any of our samples.

\textbf{4. Sequencing Library Preparation}

Once we established the successful amplification of the V3-V4 regions of
the 16S rRNA gene from our DNA extracts using an agarose gel (see
above), we further confirmed that our method successfully extracted
microbial DNA by sequencing these regions from a small subset of our
samples. Specifically, we sent seven PCR samples from two chickadee
species (\emph{Poecile atricapillus} and \emph{P. carolinensis)} plus
the 16S rRNA primers described above to Rush Genomics and Microbiome
Core Facility (Chicago, Illinois, USA) for sequencing. Library
preparation and sequencing of the seven samples were completed by the
sequencing facility using the CS1 (\emph{ACACTGACGACATGGTTCTACA}
\textbf{CCTACGGGNGGCWGCAG)} and CS2 (\emph{TACGGT-AGCAGAGACTTGGTCT}
\textbf{GACTCHVGGGTATCTAATCC)} linkers, indicated by underlining, on the
341F and 806R 16S primers, indicated in bold. The sequencing facility
performed Fluidigm amplicon library preparation to ready the samples for
next generation sequencing. The samples were normalized, pooled, and
sequenced on Illumina MiniSeq using paired end 300 bp reads. With each
run, a negative and positive controls were run alongside our samples to
control for contaminants at different stages of the sequencing.

\textbf{5. Sequencing Data Analysis}

We adapted a previously published pipeline from the R package `phyloseq'
for the statistical analysis of our microbial sequence data. Once the
sequences were obtained from the sequencing facility, we used the
program FastQC to check the initial quality of the sequenced samples and
to trim primer sequences from the samples. The quality scores indicated
the amount of overlap to use for the merged samples. To keep a minimum
phred score of 25, we merged the forward sample at 275 bp and cut the
reverse at 225 bp. The phred score is a score developed to determine the
quality of nucleobases returned from sequenced DNA. The similarity
cutoff threshold was 99\% for this sequenced data. We used the
SILVA~v138 reference taxonomy dataset to identify the microbial species
present within each sample's microbial community. Our R script is
available on GitHub
(https://github.com/rusty-russ/Russell-et-al.-Methods-Paper).

\emph{Results}

Our extraction method was successful in extracting microbial DNA from
both fecal and preen oil samples collected across a diverse range of
avian taxa (e.g., songbirds, waterfowl, scavengers, and birds of prey),
covering a broad range of feeding guilds (Tables 1 and 2; Figure 1). The
success of our extraction method was evident by our consistent ability
to PCR amplify the V3-V4 region of the microbial 16S rRNA gene,
indicated by the presence of bands on agarose gels representing PCR
products of the expected length (Figure 1). Microbial DNA was only
extracted once from each preen and fecal sample; however, multiple PCRs
were occasionally necessary (Figure 1) to visibly see bands on a 2\%
agarose gel indicating the presence of the microbial DNA. Out of 25
fecal collections from 15 species and 12 preen oil collections from 12
species, only the extraction from the single song sparrow preen oil
sample did not produce a band on an agarose gel (Figure 1).

Our successful amplification of the targeted 16S rRNA microbial gene
regions was further confirmed by sequencing a subset of our PCR
products. Specifically, we sequenced 16S rRNA V3-V4 regions in microbial
DNA extractions done with fecal material from black-capped and Carolina
chickadees (n=7). We obtained consistently high-quality reads with a
mean of 108,000 reads per sample. We obtained an initial count of 417
ASVs. After trimming and merging the reads, five of seven samples had at
least 90\% reads retained, while the remaining two had approximately
85\% reads retained (see supplemental). Additionally, after trimming and
merging and getting rid of contaminants, there remained a count of 244
ASVs. Subsequent bioinformatic analysis of the sequencing results
revealed that our extraction method was successful in extracting both
gram-positive and gram-negative bacteria (Table 3). Further, our results
identifying the bacterial classes present in each extraction, as well as
the relative abundance of different bacterial classes, suggests that the
gut microbiome can vary across different chickadee individuals (Figure
2).

\emph{Discussion}

Our new microbial DNA extraction method is effective for avian fecal
samples (Figure 1), which have been traditionally challenging materials
for use in DNA extractions. Our method is also effective for preen gland
oil samples (Figure 1), which likely harbor lower bacterial abundance
due to antimicrobial characteristics of preen oil. Both sample types
were collected across a broad range of avian species. Further, our
method successfully extracted DNA from both gram-positive and
gram-negative bacteria (Table 4). Other studies have found both gram
positive and negative bacteria, supporting the efficacy of this method.
The ease and relative cost effectiveness of our method (Table 1) makes
it particularly advantageous. We hope that our method will facilitate
advances in our understanding of microbiomes from multiple reservoirs
across a wide range of avian species.

While an organism's host microbiome is a crucial determinant of its
overall health, the full extent of the coevolutionary relationships
between a host and its symbiotic microbes is not known. Recent
discoveries have shown hosts rely on microbes for health and protection
against viruses and pathogens, providing enzymes that break down
molecules. Additionally hosts, rely on microbes for digestion, cognitive
function, growth, development, and more. Gut microbiomes are
particularly important in many taxa. To sample the gut microbiome of
birds, cloacal swabs have been shown to be nonrepresentative of
microbial diversity, only accounting for one component of the digestive
tract, whereas fecal samples contain inputs from the whole
gastrointestinal tract and are thus a more accurate representations of
the entire gut microbiome. However, existing DNA extraction methods have
not been consistently successful when working with avian fecal samples.
Birds and reptiles combine their uric waste with their fecal waste, thus
creating added difficulty in extracting DNA from fecal samples. Our
method is not only reliable with such samples but is considerably
cheaper than most commercial kits used to extract avian microbiome DNA
(Table 1). The low cost and high reliability of our method will improve
the feasibility and accessibility of research on avian host microbiomes.

As microbiome studies have increased in popularity over the last decade,
the range of host reservoirs scientists are studying has expanded as
well. Beyond the abundant and diverse gut microbiome, a potentially
functionally important reservoir in avian species is the microbiome that
exists within the uropygial or preen gland. The preen gland and the oils
produced there are known to affect communication, species recognition,
mate choice, and feather maintenance. The chemical composition of preen
oil is at least partially dependent on the microbes present in this
gland. The bacterial communities present in the uropygial oil are
thought to be less abundant than those within the gut microbiome; yet,
they are predicted to play direct roles in the odor of avian hosts as
well as potentially in feather and body health. Some preliminary studies
in this field of research have observed that disease may not be directly
linked to differences in preen oil microbiomes. Additionally,
differences in microbial communities may correspond with population
differences, and in some species, sex differences have been observed.
The consistent success of our method in extracting microbial DNA from
avian preen oil samples will facilitate new research on this currently
understudied microbiome reservoir.

Because we have established the consistent success of our method in
extracting microbial DNA from two avian sample types that are quite
chemically different, it is likely that our method will be useful across
additional avian microbiome reservoirs. Current variation in the
approaches used for microbial DNA extraction and subsequent
amplification means there is also sometimes variation in the bacteria
that are detected. Using the same extraction methodology across samples
from different reservoirs will allow more consistent and confident
comparisons of microbiomes from different reservoirs for future research
efforts. With increased interest in the relationship between hosts and
their bacterial community composition and abundance across different
microbiome reservoirs, the accurate identification of microbial taxa is
critical. With an enhanced potential to identify and quantify the
symbiotic microbes, the investigation of their impact on host health and
conservation can involve deeper questions. Widespread adoption of our
method would therefore increase the validity of future cross-study and
cross-reservoir comparisons.

To conclude, we have developed and tested a highly efficient microbial
extraction method and verified its efficacy across multiple avian
species. Our method was successful in extracting microbial DNA on the
first attempt from 25 of 25 fecal samples collected from 15 avian
species, and from 11 of 12 preen oil samples collected from 12 avian
species (Figure 1, Tables 2-3). It is also relatively inexpensive
compared with commercial fecal DNA extraction kits (Table 1). We are
confident that our method will be effective and efficient across many
additional avian species globally. Future work should explore the
effectiveness of this method in extracting microbial DNA from reservoirs
in other taxa, including reptile fecal samples as well.

\emph{Acknowledgements}

This research was supported by the U.S. National Science Foundation (NSF
IOS-1754909 and DEB-1928802 to A.M.R.) and Lehigh University Thorne
Fellowship (to A.C.R.). We thank Luke McCartin, Nicole Pittoors and
Samuel Vohsen for their assistance with microbial sequencing and
bioinformatics; and we thank Joan Spinelli, Noel Martinez, Ray Kennedy,
and Imani Ashman for providing feedback on an earlier draft of this
paper. We also thank Brandon Swayser and Haley D'Agostino at the
Wildlands Conservancy for collecting fecal samples.

\emph{}

\emph{References}

\emph{Data Accessibility Statement}

\textbf{Genetic data:}

Raw sequence reads are deposited in the NCBI Nucleotide Database
(PRJNA1015692)

\textbf{Sample metadata:}

Metadata are also stored in NCBI Nucleotide Database (PRJNA1015692)

\emph{Benefit-Sharing Statement}

Benefits Generated: Benefits from this research accrue from the sharing
of our data and results on public databases as described above.

\emph{Conflict of Interest}

The authors of this paper have no conflicts of interest.

\emph{Author Contributions}

ACR designed the experiment, optimized the method, collected, and
analyzed data, and wrote and edited the manuscript. MAK optimized the
method and edited the manuscript. AVH collected data and edited the
manuscript. AMR designed the experiment, edited the manuscript and
secured the funding.\emph{}

\emph{Tables and Figures}

\textbf{Table 1} : Cost comparison of three commercial fecal DNA
extraction kits with our extraction method (Russell et al.).\selectlanguage{english}
\begin{longtable}[]{@{}lllll@{}}
\toprule
\textbf{Extraction Method} & \textbf{Source of Reagents/Kit} &
\textbf{Cost} & \textbf{\# of Extractions} & \textbf{Cost per
extraction}\tabularnewline
\midrule
\endhead
\textbf{MP Biomedicals FastDNA-96 Fecal DNA Extraction Kit} & Fisher
Scientific & \$1,426 & 192 & \$7.43\tabularnewline
\textbf{QIAamp PowerFecal Pro~DNA Kit} & Qiagen & \$451 & 50 &
\$9.02\tabularnewline
\textbf{Quick-DNA Fecal/Soil Microbe Miniprep Kit} & Zymo-Research &
\$249 & 50 & \$4.98\tabularnewline
\textbf{Russell et al. Extraction Method} & ThermoFisher Scientific and
Fisher Scientific & \$320 & 100 & \$3.20\tabularnewline
\bottomrule
\end{longtable}

\textbf{Table 2} : Avian taxa from which we obtained fecal samples.\selectlanguage{english}
\begin{longtable}[]{@{}llllllll@{}}
\toprule
\textbf{Class} & \textbf{Order} & \textbf{Family} & \textbf{Genus} &
\textbf{Species} & \textbf{Common Name} & \textbf{Feeding Guild} &
\textbf{\# of Samples}\tabularnewline
\midrule
\endhead
Aves & Anseriformes & Anatidae & Branta & \emph{Branta canadensis} &
Canada goose & Granivore, Graminivore & 2\tabularnewline
Aves & Columbiformes & Columbidae & Columba & \emph{Columba livia} &
Rock pigeon & Granivore & 2\tabularnewline
Aves & Accipitriformes & Cathartidae & Cathartes & \emph{Cathartes aura}
& Turkey vulture & Scavenger & 1\tabularnewline
Aves & Accipitriformes & Accipitridae & Buteo & \emph{Buteo jamaicensis}
& Red-tailed hawk & Carnivore & 1\tabularnewline
Aves & Piciformes & Picidae & Dryobates & \emph{Picoides pubescens} &
Downy woodpecker & Insectivore & 1\tabularnewline
Aves & Strigiformes & Strigidae & Megascops & \emph{Megascops asio} &
Eastern screech owl & Carnivore & 1\tabularnewline
Aves & Psittaciformes & Psittaculidae & Psittacula & \emph{Melopsittacus
undulatus} & Budgerigar & Frugivore, Granivore & 2\tabularnewline
Aves & Passeriformes & Corvidae & Corvus & \emph{Corvus brachyrhynchos}
& American crow & Omnivore & 1\tabularnewline
Aves & Passeriformes & Paridae & Baeolophus & \emph{Baeolophus bicolor}
& Tufted titmouse & Insectivore & 2\tabularnewline
Aves & Passeriformes & Paridae & Poecile & \emph{Poecile carolinensis} &
Carolina chickadee & Insectivore, Frugivore & 4\tabularnewline
Aves & Passeriformes & Paridae & Poecile & \emph{Poecile atricapillus} &
Black-capped chickadee & Insectivore, Frugivore & 3\tabularnewline
Aves & Passeriformes & Sittidae & Sitta & \emph{Sitta carolinensis} &
White breasted nuthatch & Insectivore & 1\tabularnewline
Aves & Passeriformes & Icteridae & Molothrus & \emph{Molothrus ater} &
Brown-headed cowbird & Granivore & 1\tabularnewline
Aves & Passeriformes & Passerellidae & Junco & \emph{Junco hyemalis} &
Dark-eyed junco & Granivore & 2\tabularnewline
Aves & Passeriformes & Passerellidae & Melospiza & \emph{Melospiza
melodia} & Song sparrow & Insectivore & 1\tabularnewline
\bottomrule
\end{longtable}

\textbf{Table 3} : Avian taxa from which we obtained preen oil samples.\selectlanguage{english}
\begin{longtable}[]{@{}lllll@{}}
\toprule
\textbf{Order} & \textbf{Family} & \textbf{Genus} & \textbf{Species} &
\textbf{Common Name}\tabularnewline
\midrule
\endhead
Passeriformes & Corvidae & Corvus & \emph{Corvus brachyrhynchos} &
American crow\tabularnewline
Passeriformes & Paridae & Poecile & \emph{Poecile carolinensis} &
Carolina chickadee\tabularnewline
Passeriformes & Paridae & Poecile & \emph{Poecile atricapillus} &
Black-capped chickadee\tabularnewline
Passeriformes & Turdidae & Turdus & \emph{Turdus migratorius} & American
robin\tabularnewline
Passeriformes & Sittidae & Sitta & \emph{Sitta carolinensis} &
White-breasted nuthatch\tabularnewline
Passeriformes & Fringillidae & Carduelis & \emph{Carduelis tristis} &
American goldfinch\tabularnewline
Passeriformes & Fringillidae & Carpodacus & \emph{Carpodacus mexicanus}
& House finch\tabularnewline
Passeriformes & Cardinalidae & Cardinalis & \emph{Cardinalis cardinalis}
& Northern cardinal\tabularnewline
Passeriformes & Passerellidae & Zonotrichia & \emph{Zonotrichia
albicollis} & White-throated sparrow\tabularnewline
Passeriformes & Passerellidae & Melospiza & \emph{Melospiza melodia} &
Song sparrow\tabularnewline
Passeriformes & Icteridae & Agelaius & \emph{Agelaius phoeniceus} &
Red-winged blackbird\tabularnewline
Passeriformes & Icteridae & Molothrus & \emph{Molothrus ater} &
Brown-headed cowbird\tabularnewline
\bottomrule
\end{longtable}

\textbf{Table 4} : Gram-positive and gram-negative categorization of the
bacterial taxa detected from the amplification of the 16S rRNA gene from
seven microbial DNA samples extracted using our method.\selectlanguage{english}
\begin{longtable}[]{@{}ll@{}}
\toprule
\textbf{Gram-Positive} & \textbf{Gram-Negative}\tabularnewline
\midrule
\endhead
Acidimicrobiia & Acidobacteriae\tabularnewline
Alphaproteobacteria & Alphaproteobacteria\tabularnewline
Actinobacteria & Bacteroidota\tabularnewline
Firmicutes & Bdellovibrionota\tabularnewline
& Chloroflexi\tabularnewline
& Cyanobacteriia\tabularnewline
& Firmicutes\tabularnewline
& Fusobacteriota\tabularnewline
& Gemmatimonadota\tabularnewline
& Planctomycetota\tabularnewline
& Pseudomonadota\tabularnewline
& Verrucomicrobia\tabularnewline
\bottomrule
\end{longtable}\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image1/image1}

\end{center}
\end{figure}

\textbf{Figure 1} : Phylogenetic relationships of the avian taxa from
which we obtained fecal and preen samples. The phylogenetic tree was
made using Vertlife.org. The circle indicates a preen oil sample was
collected from that species and the square indicates a fecal sample was
collected from that species. The number of PCR rounds run on the
original extracted DNA to successfully see a band on a 2\% agarose gel
is indicated by the color and pattern within each circle or square. The
V3-V4 region of the microbial 16S rRNA gene successfully amplified from
all but one of the DNA extracts we generated using our method.\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image2/image2}

\end{center}
\end{figure}

\textbf{Figure 2} : Relative abundance of several bacterial classes
present in the microbial DNA extracted from three black-capped
(BC;\emph{Poecile atricapillus} ) and four Carolina chickadee (CA;
\emph{P. carolinensis} ) fecal samples. See text for an explanation of
how the microbial species present were identified using sequence data.

\selectlanguage{english}
\FloatBarrier
\end{document}

