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\begin{document}

\title{Genomic features underlying evolutionary transitions of Apibacter to
honeybee gut symbionts}



\author[1]{Wenjun Zhang}%
\author[1]{Xue Zhang}%
\author[1]{Qinzhi Su}%
\author[1]{Min Tang}%
\author[1]{Hao Zheng}%
\author[2]{Xin Zhou}%
\affil[1]{China Agricultural University}%
\affil[2]{Beijing Advanced Innovation Center for Food Nutrition and Human Health, China Agricultural University, Beijing 100193, People’s Republic of China}%


\vspace{-1em}



  
  \date{June 18, 2024}


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\selectlanguage{english}
\begin{abstract}
The symbiotic bacteria associated with honeybee gut have likely
transformed from a free-living or parasitic lifestyle, through a close
evolutionary association with the insect host. However, little is known
about the genomic mechanism underlying bacterial transition to exclusive
adaptation to the bee gut. Here we compared the genomes of bee gut
symbionts Apibacter with their close relatives living in different
lifestyles. We found that despite of general reduction in the Apibacter
genome, genes involved in amino acid synthesis and monosaccharide
detoxification were retained, which was likely beneficial to the host.
Interestingly, the microaerobic Apibacter species have specifically
preserved the NAR operon encoding for the nitrate respiration pathway
which in contrast, is absent from the related non-free-living
microaerobic pathogenic relatives. The NAR operon is also conserved in
the cohabiting bee microbe Snodgrasella, but with a differed structure.
This convergence implies a crucial role of respiration nitrate reduction
for microaerophilic microbiomes to colonize bee gut epithelium. Genes
involved in lipid, histidine and phenylacetate degradation are partially
lost in Apibacter, possibly associated with the loss of pathogenicity.
Antibiotic resistance genes were only sporadically distributed among
Apibacter species, but condensed in their pathogenic relatives.
Collectively, this study advanced our understanding of genomic
transition underlying specialization in bee gut symbionts.%
\end{abstract}%



\sloppy




\section*{Introduction}

{\label{introduction}}

Bacterial symbiotic association with insect host is ubiquitous in
nature, which confers the host with traits to explore new ecological
niches (Sudakaran, Kost, \& Kaltenpoth, 2017). Symbiotic bacteria
inhabit either intra- or extra-cellularly inside the insect, providing
hosts with vital benefits, including nutrition supply, pathogen
resistance, assistance in immunity development, among others (Engel \&
Moran, 2013; McCutcheon, Boyd, \& Dale, 2019). Mutualistic symbionts may
have various origins, including environmental bacteria or infective
parasites (Sachs, Skophammer, \& Regus, 2011; Sachs, Skophammer, Bansal,
\& Stajich, 2013). Despite of distinct evolutionary pathways, the
convergent transition to a mutualistic symbiotic lifestyle often
involves the loss of virulence factors, degeneration of superfluous
functions, and preservation or obtaining of traits beneficial to the
host (EWALD, 1987; McCutcheon \& Moran, 2012). However, direct evidence
showing taxonomic turn-over in insect symbionts are scarce.
Alternatively, comparative genomic analysis of mutualistic symbionts
with the free-living lineages that are phylogenetically closely related
provides a feasible route to examine the evolutionary pattern and
adaptive mechanism accompanying the evolutionary transition to mutualism
(Boscaro et al., 2013; Zheng, Dietrich, \& Brune, 2017).

The honeybees have simple but specific gut symbiotic bacteria from
genera of \emph{Gilliamella} , \emph{Snodgrasella} ,
\emph{Lactobacillus}and \emph{Bifidobacterium} (Martinson, Moy, \&
Moran, 2012). They compose more than 95\% of the whole gut community,
and the association with these five core bacteria can be dated back
prior to the divergence of corbiculate bees (i.e., honeybee, bumblebee,
stingless bee, and orchid bee) (Kwong et al., 2017). Although bee gut
bacteria are not intracellular inhabitant and maternally transferred,
they are inheritable among worker bees in the colony through social
contacts (Powell, Martinson, Urban-Mead, \& Moran, 2014). Along with the
establishment of symbiotic association, these bacteria are subjected to
genome reduction and evolutionary adaptation (Kwong, Engel, Koch, \&
Moran, 2014). This adaptive transition increases their fitness to the
bee gut niche, but also limits the capacity to thrive in other
environments (Ellegaard et al., 2019). As with other symbiotic microbes,
honeybee gut bacteria share ancestry with bacterial species carrying
varied lifestyles (Segers, Ke\selectlanguage{polish}š\selectlanguage{english}nerov\selectlanguage{ngerman}á, Kosoy, \& Engel, 2017).
Subsequently, genomic changes are expected to be remarkably different
between the transitions from the common ancestry to gut symbionts and to
other lifestyles (Tamarit et al., 2015). Thus, the honeybee-gut bacteria
system provides a promising model to explore the genomic features
underlying lifestyle transition of free-living bacteria to mutualistic
gut symbionts.

\emph{Apibacter} is a genus of bacteria that is prevalent in \emph{Apis
cerana} , \emph{Apis dorsata} and bumblebee species (genus\emph{Bombus}
), however, they are only sporadically reported in\emph{Apis mellifera}
(Kwong et al., 2017; Kwong \& Moran, 2016). Our knowledge on
\emph{Apibacter} is primarily based on genome sequences of only four
strains till now (Kwong, Steele, \& Moran, 2018). Phylogenetic analysis
showed that \emph{Apibacter} spp. form a monophyletic lineage, which is
embedded in the \emph{Chryseobacterium} clade
(family\emph{Flavobacteriaceae} ) (Kwong \& Moran, 2016). The members of
the\emph{Chryseobacterium} clade show a variety of lifestyles, including
environmental free-living, opportunistic or obligate mammal and bird
pathogens (McBride, 2014), of which the genus \emph{Chryseobacterium} is
the dominant member and consist of mostly environmental habitants. But
strains of \emph{Chryseobacterium gleum} and \emph{Chryseobacterium
indologenes} are common clinical isolates, which cause infections in
immunocompromised patients (Calderán et al., 2011). Type strains of
species from \emph{Elizabethkingia} and \emph{Empedobacter} are also
opportunistic human pathogens that infect urinary and respiratory tract,
causing sepsis, pneumonia and meningitis (Bhat, Priya, Krishnan, \&
Kanungo, 2016; Gupta, Zaman, Mohan, \& Taneja, 2017; Lau et al., 2015;
Zaman, Gupta, Kaur, Mohan, \& Taneja, 2017). Isolates of
genera\emph{Bergeyella} and \emph{Weeksella} infect mammal respiratory
tracts and female genitourinary tract, respectively (Hugo, Bruun, \&
Jooste, 2006; Lin, Chen, \& Liu, 2007; Slenker, Hess, Jungkind, \&
DeSimone, 2012). Members of \emph{Riemerella} and
\emph{Ornithobacterium} are important bird pathogens, causing infections
to respiratory tract and other organs (McBride, 2014). These bacteria
can be transmitted horizontally by aerosol or vertically between host
generations. Type species of genera \emph{Bergeyella} , \emph{Weeksella,
Riemerella} and\emph{Ornithobacterium} are considered non-free-living,
showing a preference to micro-aerophilic conditions, which is different
from most other strictly aerobic members of the \emph{Chryseobacterium}
clade (Mavromatis et al., 2011; Van Empel \& Hafez, 1999). The common
features of the \emph{Chryseobacterium} clade include proteolytic
activity, carbohydrate utilization and multiple antibiotic resistance,
causing challenges to the treatment of infection. Protease and biofilm
formation are suggested to be important for their pathogenesis (McBride,
2014). However, the genetic mechanisms underlying the pathogenicity
remain uncovered.

In this study, we isolated and sequenced the genomes of
14\emph{Apibacter} strains from the Asian honeybee \emph{Apis cerana} .
Molecular and physiology experiments were conducted to characterize its
colonization positions and specificity inside the bee gut. Comparative
genomic analyses of honeybee symbiotic \emph{Apibacter} spp. with
phylogenetically related free-living and pathogenic strains from
the\emph{Flavobacteriaceae} family revealed key genomic changes
of\emph{Apibacter} underlying adaptation to mutualistic symbiotic
relationship with the honeybee.

\section*{Materials and Methods}

{\label{materials-and-methods}}

\subsection*{\texorpdfstring{Sample collection and
\emph{Apibacter}isolation}{Sample collection and Apibacterisolation}}

{\label{sample-collection-and-apibacter-isolation}}

The worker bees of \emph{Apis cerana} were collected from Sichuan, Jilin
and Qinghai Provinces in China (Dataset S1). A total of six guts were
dissected, homogenized and frozen in glycerol (25\%, vol/vol) at --80°C.
Frozen stocks of homogenized guts were streaked out on the heart
infusion agar (Oxoid) or Columbia agar (Oxoid) supplemented with 5\%
sheep blood (Solarbio). The plates were incubated at 35°C in 5 \%
CO\textsubscript{2} for 2--3 days. Colonies were screened by PCR using
universal primers 27F and 1492R for the16S rRNA gene and Sanger
sequencing.

\subsection*{\texorpdfstring{Fluorescence \emph{in situ} hybridization
(FISH)
microscopy}{Fluorescence in situ hybridization (FISH) microscopy}}

{\label{fluorescence-in-situ-hybridization-fish-microscopy}}

Ten adult worker bees of \emph{A. cerana} were collected from a single
colony in Beijing, China in May 2018. The whole guts of bees were
dissected and frozen in RNAlater (Qiagen) at -80 ºC. The FISH protocol
was adapted from Yuval et al. (Gottlieb et al., 2006). In brief, the
guts preserved in RNAlater were fixed in the fixative solution (Ethanol:
Chloroform: Acetic acid = 6:3:1) and kept overnight at the room
temperature, followed by rinsing with 1 × PBS. The guts were then
treated with 1 mg/mL proteinase K solution at 56 °C for 20 minutes,
followed by rinsing with 1 × PBS. The guts were incubated with 6\%
H\textsubscript{2}O\textsubscript{2} ethanol solution for 2 hours,
followed by rinsing twice with 1 × PBS. Finally, the guts were permeated
with 0.1\% Triton for 2 hours, followed by rinsing for 2 or 3 times in 1
× PBS buffer.

Binding FISH probes targeting 16S rRNA genes were designed specifically
for each of the genera \emph{Apibacter} , \emph{Snodgrasella}
and\emph{Gilliamella} , following Martinson et al. (Martinson et al.,
2012) (Table S1). Probe hybridization was performed overnight at 37 °C,
followed by rinsing in 1 × PBS for 3 times. Spectral imaging was used to
visualize gut sections on a ZEISS LSM780 confocal microscope.
Autofluorescence was assayed for each tissue type as the negative
control. \emph{Apibacter} were labeled with florescent together with
either \emph{S. alvi} or \emph{G. apicola} .

\subsection*{Estimation of bacterial abundance using
qPCR}

{\label{estimation-of-bacterial-abundance-using-qpcr}}

Worker bees were collected from 2 colonies at the same apiary in
Beijing, China in September 2018. Samples were stored at -80°C. Gut
segments (i.e., midgut, ileum, and rectum) were dissected and separated.
DNA was extracted following Powell et al. (Powell et al., 2014).
Universal 16S primers 27F and 355R for bacteria were employed
and\emph{Apibacter} -specific primers Apiq9-F and Apiq9-R were designed
in this study (Table S1). Standard curves were created and reactions
were carried out on a LightCycler 480 (Roche Applied Science,
Indianapolis, IN). Significance in differences within and between
samples was determined using Mann-Whitney (Wilcoxon-rank) nonparametric
U tests.

\subsection*{\texorpdfstring{\emph{In vivo} colonization
experiments}{In vivo colonization experiments}}

{\label{in-vivo-colonization-experiments}}

Microbiota-depleted bees were obtained following Zheng et al. (Zheng,
Powell, et al., 2017). Late stage pupae of both \emph{A. cerana}
or\emph{A. mellifera} were removed from brood frames and incubated for
24-36 h in sterile plastic bins at 35 °C and 75\% humidity. Newly
emerged bees were kept in cup cages provided with sterilized sucrose
syrup (0.5 M) and bee bread. For inoculation, bacteria strain of
the\emph{Apibacter} sp. B3706 was grown on the heart infusion agar
(Oxoid), which was supplemented with 5\% sheep blood from glycerol
stocks, for two days. Cultivated strains were scraped and suspended in 1
× PBS to reach an OD\textsubscript{600} of 1.0. Batches of 25 bees were
placed in a 50 ml conical tube, and 50 \selectlanguage{greek}μ\selectlanguage{english}l sucrose syrup was added. The
tube was rotated gently so that the syrup was coated on the surface of
bees. The tube was rotated again after the addition of 50 \selectlanguage{greek}μ\selectlanguage{english}l of
the\emph{Apibacter} sp. B3706 suspension (\textasciitilde{}2 \selectlanguage{ngerman}×
10\textsuperscript{6} cells per bee). The bacteria were inoculated into
the bee guts as a result of auto- and allogrooming. Inoculated bees were
reared in cup cages. Three replicate enclosures were set up for
both\emph{A. cerana} or \emph{A. mellifera} with 20 bees in each cage.
The inoculated bees were fed with sucrose syrup and sterilized pollen
throughout the experiment. Colonization levels were determined at day 6
using qPCR as described previously.

\subsection*{Genome sequencing, assembly and
annotation}

{\label{genome-sequencing-assembly-and-annotation}}

Genomic DNA of \emph{Apibacter} isolates were extracted using a
bead-beating method as previously described (Powell et al., 2014). Two
strains (\emph{Apibacter} sp. B2966 and B3706) were sequenced with the
PacBio RS (PacBio) at Nextomics Biosciences Co. Ltd., China, and
assembled using OLC algorithm of the Celera (Chaisson \& Tesler, 2012).
Three strains (\emph{Apibacter} sp. B3239, B3546 and B2912) were
sequenced with a BGISEQ-500RS (BGI) at BGI-Qingdao, China. The other
nine strains (\emph{Apibacter} sp. B3813, B3883, B3887, B3889, B3912,
B3913, B3918, B3924 and B3935) were sequenced with a HiSeq X-Ten
(Illumina) at Novogene Co., Ltd, China. The twelve strains except
for\emph{Apibacter} sp. B2966 and B3706 were assembled
with\emph{SOAPdenovo-Trans} (version 2.04, -K 51 -m 91 --R for 150PE
reads; -K 31 -m 63 --R for 100PE reads) (Xie et al., 2014),
\emph{SOAPdenovo}(only for 150PE reads) and \emph{SPAdes} (version
3.13.0, -k 33,55,77,85) (Bankevich et al., 2012). The quality trimmed
reads were mapped back to the assembled contigs using minimap2-2.9 (Li,
2018) to examine assembly quality. The bam file generated by
\emph{samtools}(version 1.8) (Li et al., 2009) and the assemblies were
processed
by\emph{BamDeal}(\emph{https://github.com/BGI-shenzhen/BamDeal}, version
0.19) to calculate and visualize sequencing coverage and GC contents of
assembled contigs. Spurious contaminants, contigs with low depths,
unnormal GC contents and those apparently differing from the cluster,
were removed from the draft genome. Assembled genomes were then
annotated using PROKKA 1.13.3 (Seemann, 2014).

\subsection*{Comparisons of genome structure, genome divergence, and gene
contents}

{\label{comparisons-of-genome-structure-genome-divergence-and-gene-contents}}

The contigs of each assembly were re-ordered according to the single
circular genome of strain B3706 using the `Contig Mover' tool of Mauve
version 2.4.0 (Darling, Mau, Blattner, \& Perna, 2004; Rissman et al.,
2009). Pairwise average nucleotide identity (ANI) was calculated with
JSpeciesWS (Richter, Rosselló-Móra, Oliver Glöckner, \& Peplies, 2016)
using the BLASTN algorithm (ANIb). Genome completeness was estimated
with CheckM (Parks et al., 2015), which was available at KBase online
(Arkin, Stevens, Cottingham, Maslov, \& Perez, 2015) using recommended
parameters. The genome structures were compared using the R-package
genoPlotR (Guy, Kultima, Andersson, \& Quackenbush, 2011).

\subsection*{Phylogenetic inference}

{\label{phylogenetic-inference}}

Gene orthology was determined using OrthoMCL (Li et al., 2003) for all
genomes used in this study. All steps of the OrthoMCL pipeline were
executed as recommended in the manual and the mcl program was conducted
using parameters `-abc -I 1.5'.

Protein sequences of the identified single-copy orthologous were aligned
using Mafft-linsi (Katoh, Kuma, Miyata, \& Toh, 2005). Alignment columns
only containing gaps were removed and the alignments were concatenated.
The phylogeny was reconstructed using RAxML v8.2.10 (Stamatakis, 2014)
with the PROTGAMMAIJTT model and 100 bootstrap replicates.

\subsection*{Gene flux analysis}

{\label{gene-flux-analysis}}

Gene gain and loss analyses and inferences for gene contents of LCAs
(last common ancestors) were conducted using Count (Csurös, 2010).
Standard methods used in previous works were employed in the present
study (Segers et al., 2017). For each gene family, Wagner parsimony with
a gene gain/loss penalty of 2 (Zaremba-Niedzwiedzka et al., 2013) was
used to infer the most parsimonious ancestral states. Parameter choices
followed a previous publication (Oyserman et al., 2016).

\subsection*{Analysis of functional gene
contents}

{\label{analysis-of-functional-gene-contents}}

Gene contents were categorized based on COG and eggNOG (Huerta-Cepas et
al., 2019) functions. Significantly differential genes and pathways were
enriched by OrthoVenn (Yi, Colemanderr, Chen, \& Gu, 2015). For gene
families subset of interest, BLASTP(Altschul et al., 1997) was used to
query against the NCBI's nr database and TIGRFAM Hidden Markov Model
(HMM) (Haft, Selengut, \& White, 2003). The phylogenies of
\emph{narG}and \emph{narH} were produced from protein sequences obtained
by blastp against NCBI's nr database with default parameters. And the
hits aligned using Mafft-linsi, trimmed with trimAL (Capella-Gutiérrez,
Silla-Martínez, \& Gabaldón, 2009) for sites with over 50\% gaps and
phylogenetic trees were constructed using RAxML PROTGAMMALG with 100
bootstraps. The parameters were based on previous publication (Neuvonen
et al., 2016).

Carbohydrate-active enzyme (CAZymes) gene families were identified for
all analyzed genomes using the command-line version of dbCAN (Database
for automated Carbohydrate-active enzyme Annotation) (Yin et al., 2012),
following authors' instruction. The PULs (Polysaccharide utilization
loci) were identified using the TIGRfam (Potter et al., 2018) and Pfam
(Finn et al., 2014) models, following Terrapon et al. (Terrapon,
Lombard, Gilbert, \& Henrissat, 2015). Antibiotic resistance genes were
identified by querying all the genomes against the Comprehensive
Antibiotic Resistance Database (CARD) (McArthur et al., 2013).

\section*{Results}

{\label{results}}

\textbf{Spatial distribution of \emph{Apibacter} in the gut of \emph{A.
cerana}}

We characterized the colonization of \emph{Apibacter} in the gut
of\emph{A. cerana} using fluorescence in situ hybridization (FISH) and
qPCR. Honeybee symbionts \emph{Snodgrassella} and \emph{Gilliamella} are
dominant in the ileum, with \emph{Snodgrassella} colonizing the inner
wall of ileum (Martinson et al., 2012). These two bacteria were used as
reference coordinates to infer the location of \emph{Apibacter} using
species-specific probes (Fig. 1A-D). The spectral images showed
that\emph{Apibacter} co-resided with \emph{Snodgrassella} in both ileum
and midgut (Fig. 1A, C). The signals were stronger at the inner walls,
indicating that \emph{Apibacter} colonized the gut intima as
did\emph{Snodgrassella} . \emph{Gilliamella} covered on the top
of\emph{Apibacter} and extended into the gut lumen (Fig. 1B,
D).\emph{Apibacter} could not be visualized clearly in the rectum, as
the rectum was filled with pollen grains with auto-fluorescent under the
excitation wavelength. We also quantified the absolute abundances
of\emph{Apibacter} in different gut compartments (midgut, ileum, rectum)
using qPCR. It showed that the absolute abundances of
\emph{Apibacter}increased from midgut to rectum, with cell numbers
ranged from 9.43×10\textsuperscript{5} to 1.74×10\textsuperscript{7}
(Fig. 1E).

Host specialization is a common feature for many host-microbiome
associations (Oh et al., 2010), which has been demonstrated by the
honeybee symbiotic bacteria \emph{Snodgrassella} (Kwong et al., 2014)
and \emph{Lactobacillus} (Ellegaard et al., 2019). To test if it is also
the case for \emph{Apibacter} , we performed the colonization
specificity test. \emph{Apibacter} sp. strain B3706 isolated from
\emph{A. cerana}was inoculated to the microbiota-free \emph{A. cerana}
and \emph{A. mellifera} (see Methods). Six days after inoculation, the
cell numbers in the gut were more than 10\textsuperscript{7}, which were
much higher than in the inoculum (\textless{} 10\textsuperscript{6}),
indicating that strain B3706 was able to colonize the guts of both
\emph{A. cerana}and \emph{A. mellifera} . However, the colonization
efficiency in\emph{A. cerana} was significantly higher than that in
\emph{A. mellifera} (Fig. 1F), suggesting that strain B3706 was less
adapted to the gut of \emph{A. mellifera} .

\textbf{Genome characters of \emph{Apibacter} and phylogenetic
inference}

We isolated 14 strains of \emph{Apibacter} from worker bees of \emph{A.
cerana} from Sichuan, Jilin, and Qinghai provinces in China (Dataset
S1). Two genomes (strains B3706 and B2966) were completed into single
circular chromosomes by sequencing on the PacBio platform. The
assemblies of genomes sequenced with either Illumina or BGISEQ contained
12--49 contigs and with full completeness as evaluated by CheckM (Parks,
Imelfort, Skennerton, Hugenholtz, \& Tyson, 2015) (Table
S2).\emph{Apibacter} strains isolated from \emph{A. cerana} had genome
sizes ranging from 2.26 to 2.35 Mb, similar to those from the bumble
bees (2.33 Mbp; Brandt et al. 2016), but smaller than those from
\emph{A. dorsata} (2.63--2.76 Mbp; Kwong, Steele, and Moran 2018) (Table
S2). Parasitic pathogenic species from genera \emph{Riemerella}
,\emph{Bergeyella} , \emph{Weeksella} , and \emph{Ornithobacterium} also
have small genome sizes ranging from 2.16--2.44 Mb comparing to other
type strains in the \emph{Chryseobacterium} clade, suggesting that they
are subjected to genome reduction as documented for other symbionts
(Fig. S2 B, Table S2) (Pérez-Brocal, Latorre, \& Moya, 2013). The GC
contents of the honeybee symbiotic \emph{Apibacter} are lower than its
non-symbiotic relatives, a common feature of symbionts attributed to the
mutational bias and weak selections (Fig. S2 B) (McCutcheon et al.,
2019; McCutcheon \& Moran, 2012).

The average nucleotide identities (ANIs) between genomes obtained in
this study and those from Malaysian \emph{A. cerana}
(\emph{Apibacter}sp. wkB309, Kwong \& Moran, 2016) are 95.82\%--97.33\%,
which suggests that \emph{Apibacter} carried by the mainland \emph{A.
cerana} have diverged from those from the Sundaland \emph{A. cerana} ,
on the verge of speciation (Table S2). Six genomes among the 14
sequenced strains have almost identical ANIs (99.99\%), the consensus of
which was used in subsequent analyses. The genomic divergence in
\emph{Apibacter} isolates is more obvious between bee hosts, as the ANIs
are only 85\% and 74\% to those isolated from bumble bees and \emph{A.
dorsata} , respectively. Despite of large sequence divergence, genome
structures are mostly conserved across the \emph{Apibacter} strains from
\emph{A. cerana} and\emph{Bombus} , even though few gene rearrangements
and inversions are observed in \emph{Apibacter} genomes isolated from
\emph{A. dorsata}(Fig. S1). But the overall structural variation seems
not substantial when \emph{Apibacter} diverged with the hosts. Such a
pattern is in congruent with the observations in \emph{Bartonella}
\emph{apis} from\emph{A. mellifera} , and in \emph{Buchnera aphidicola}
from aphids (Chong, Park, \& Moran, 2019; Segers et al., 2017).

A maximum-likelihood phylogeny was inferred using a total of 681
orthologous single-copy genes present in all \emph{Apibacter,} related
taxa and one outgroup (\emph{Flavobacterium aquatile} LMG 4008) chosen
from the \emph{Flavobacteriaceae} family (Fig. S2 A) (Li, Stoeckert, \&
Roos, 2003; Stamatakis, 2014). Consistent with the previous phylogenetic
relationship (Kwong \& Moran, 2016), \emph{Apibacter} species formed a
monophyletic group and the isolates from \emph{A. cerana} clustered
together (Fig. S2 A). The \emph{Apibacter} clade was sister the lineage
consisting of \emph{Elizabethkingia} , \emph{Riemerella}
and\emph{Chryseobacterium} genera, which is referred to as Clade C
hereafter following Kwong and Moran (Kwong \& Moran, 2016). These genera
contain environmental non-pathogens, opportunistic pathogens, and
parasitic pathogens (McBride, 2014). Finally, the \emph{Apibacter} clade
and Clade C together formed a sister relationship to Clade E
(\emph{Empedobacter} , \emph{Weeksella} , and \emph{Ornithobacterium} ),
which consists of free-living and parasitic strains associated with
mammal and bird diseases (McBride, 2014).

\subsection*{\texorpdfstring{\emph{Apibacter} conferred a large number of
genes loss but preserved specific host beneficial
functions}{Apibacter conferred a large number of genes loss but preserved specific host beneficial functions}}

{\label{apibacter-conferred-a-large-number-of-genes-loss-but-preserved-specific-host-beneficial-functions}}

To infer the genes loss patterns in \emph{Apibacter} spp. during
adaptive transition to a bee gut symbiont, we constructed the last
common ancestor (LCA) for \emph{Apibacter} spp. and close relatives
using generalized parsimony and analyzed the gene flux. A total of 601
genes were lost at the node leading to the LCA of \emph{Apibacter}branch
(Fig. 2A). Of these, 498 were preserved in both Clade C and E, and 99
were only present in Clade C. Unique genes present in Clade C but lost
in \emph{Apibacter} might be associated with non-symbiotic lifestyles,
e.g. environmental free-living. Among genes with known functions, those
lost in \emph{Apibacter} are enriched in the COG categories of inorganic
ion transport, transcription, amino acid transport and metabolism,
carbohydrate transport and metabolism, cell wall biogenesis and lipid
transport and metabolism (8.7, 8.5, 8.1, 7.7, 7.7 and 6.5\% of genes
with COG annotation respectively, Dataset S2). The loss of cell wall
biosynthesis is in line with the general pattern of genome reduction
found in symbionts, which facilitates the exchange of molecules through
the host-symbionts interface and allows further degeneration of
metabolites in the symbionts (McCutcheon \& Moran, 2012). The losses of
amino acid membrane transport, carbohydrate and lipid metabolic
processes could be explained as an adaptation to a nutrient-rich
environment (Schmid et al., 2018).

It showed that a few symbiotic bacteria could provide major functions in
pollen degradation in the bee gut, facilitating polysaccharide
utilization for the host (Ke\selectlanguage{polish}š\selectlanguage{english}nerov\selectlanguage{ngerman}á et al., 2017), similar
to\emph{Bacteroidates} in human gut (Koropatkin, Cameron, \& Martens,
2012). However, genes involved in carbohydrate metabolism are
substantially lost in \emph{Apibacter} . Compared with Clade C and
E,\emph{Apibacter} lost most of the carbohydrate active enzymes
(CAZymes) genes and retained no more than two polysaccharide utilization
loci (PUL). The remaining PULs in \emph{Apibacter} are composed of only
tandem \emph{susCD} -like genes, and lack all surrounding CAZyme genes
(Dataset S3). The protein sequences of these PUL residue structures are
conserved across \emph{Apibacter} isolates from \emph{A. cerana}
and\emph{Bombus} (\textgreater{}87\% similarity). To the contrast, the
PULs in \emph{Apibacter} genomes from \emph{A. dorsata} are more
diverse, with one from wkB301 homologous to those from \emph{A. cerana}
and bumble bee (\textgreater{}70\% similarity), one conserved among
wkB301 and wkB180 from \emph{A. dorsata} (\textgreater{}95\%
similarity), and one unique to wkB180 (isolate of bumble bee, Dataset
S3). The variations in PUL residue structure among \emph{Apibacter}
strains suggests independent gene losses of CAZyme genes in the \emph{A.
dorsata}lineage.

In contrast to significant gene loss in \emph{Apibacter} , specific
genes beneficial to the host were preserved. For instance, the
mannose-6-phosphate isomerase encoded by \emph{manA} is responsible for
degradation of the toxic mannose for the host (Zheng et al., 2016). This
gene is lost in all strains of Clade C and E, but is preserved by
all\emph{Apibacter} strains, indicating that mannose detoxification is
an important mutualistic trait in \emph{Apibacter} (Dataset S5). It is
known that symbiotic gut bacteria are capable of synthesizing amino
acids for host and other co-occuring symbionts (Kwong et al., 2014;
McCutcheon \& Moran, 2012). Consistently, \emph{Apibacter} have
preserved all genes underlying amino acid synthesis, which are inherited
from the LCA. On the contrary, the parasitic pathogens from
genera\emph{Bergeyella} , \emph{Weeksella} , \emph{Riemerella}
and\emph{Ornithobacterium} , have all conferred a genome reduction,
while losing substantial amino acid synthesis genes (Dataset S4)
(Rohmer, Hocquet, \& Miller, 2011).

\subsection*{\texorpdfstring{Respiratory nitrate reduction was enriched
and conservative
in\emph{Apibacter}}{Respiratory nitrate reduction was enriched and conservative inApibacter}}

{\label{respiratory-nitrate-reduction-was-enriched-and-conservative-in-apibacter}}

A total of 1,349 gene families are shared by all three clades,
representing core gene functions shared by bacteria of varied lifestyles
(Fig. 2C). \emph{Apibacter} LCA has 226 unique gene families, which
include functions that are specific to the adaptation to the bee host.
Clade C and Clade E have more gene families (467 gene families) in
common compared to what is shared with the \emph{Apibacter} group, in
congruent with the non-symbiotic lifestyles shared between them. Gene
ontology (GO) enrichment analysis was performed to explore
overrepresented functions specific to the \emph{Apibacter} group.
Interestingly, the gene families belonging to the respiratory nitrate
reduction (NAR) pathway are enriched in \emph{Apibacter} (Dataset S5).
Furthermore, the biosynthesis of the molybdenum cofactors (Moco), which
are required for NAR nitrate reductase (Moreno-Vivián, Cabello,
Martínez-Luque, Blasco, \& Castillo, 1999; Stewart, 1988), are also
enriched. These genes locate in the NAR operon and are harbored in
all\emph{Apibacter} isolates, with only one Moco synthesis gene (mosC)
and one nitrate transporter missing in two \emph{A. dorsata} isolates
(Fig. 3). The NAR related genes are highly conserved in amino acid
sequences among genomes isolated from the same bee hosts
(\textgreater{}94\% similarity). Isolates from \emph{A. cerana} are more
similar to the one from bumble bee than those from \emph{A. dorsata}
(Fig. 3). The function of the NAR pathway seems to be highly conserved,
eventhough a key member gene is replaced by alternative gene. The
\emph{narI} encoding the membrane biheme b quinol-oxidizing \selectlanguage{greek}γ \selectlanguage{english}subunit of
the nitrate reductase is missing. Alternatively, a Rieske protein
homolog was identified in the NAR operon. The Rieske protein is an
iron-sulfur protein (2Fe-2S), with a function in transferring electrons
from the quinone pool, which is equivalent to NarI (Schneider \&
Schmidt, 2005). Therefore, the Rieske protein homolog is expected to
have replaced the function of NarI in transferring electrons to the
NarGH complex (Arshad et al., 2015). The replacement of \emph{narI} in
the NAR operon structure was previously identified in halophilic
archaea, which represents an ancient respiratory nitrate reductase
(Cabello, Rold\selectlanguage{ngerman}án, \& Moreno-Vivián, 2004; Yoshimatsu, Iwasaki, \&
Fujiwara, 2002). Additionally, three copies of the \emph{narK} nitrate
transporter gene are identified in\emph{Apibacter} (Fig. 3B) (Cole \&
Richardson, 2017), implying high efficiency of nitrate respiration in
\emph{Apibacter} . To infer the evolutionary origin of the nitrate
reductase genes, the Maximum likelihood tree based on the \emph{narG}
and \emph{narH} genes were constructed (Fig. S4). Interestingly, the
\emph{narG} and \emph{narH} of\emph{Apibacter} formed a monophyletic
clade. The phylogenetic relationship of \emph{narG} and \emph{narH} is
consistent with the phylogeny of the bacterial strains that harbor the
genes. Their closely related genes are mostly from strains of genera
\emph{Flavobacterium} , implying that the nitrate reductase genes of
\emph{Apibacter} are vertically inherited.

In contrast, bacteria from Clade C and E lack intact NAR pathways, which
is congruent with their aerobic nature. It is worth noting that
parasitic pathogens from \emph{Riemerella} and
\emph{Ornithobacterium}also lack the NAR operon ( Mavromatis et al.,
2011; Van Empel \& Hafez, 1999), despite that they are microaerophilic,
as with \emph{Apibacter}(Fig. 3, Dataset S5). These results imply that
the NAR pathway might be particularly beneficial to gut commensal
bacteria, which prompted us to survey the NAR pathway in other honeybee
gut bacteria. Interestingly, NAR pathway is also mostly conserved in
\emph{Snodgrassella} strains, but absent in the other four core bee gut
bacterial phylotypes (\emph{Gilliamella} , \emph{Bifidobacterium} ,
\emph{Lactobacillus} Firm4 and Firm5). This difference further suggests
that the NAR pathway might be generally required by microaerobic
bacteria inhabiting gut epithelium (Dataset S5). However, the nitrate
reductase of \emph{Snodgrassella}showed obvious variations when compared
with that of \emph{Apibacter} : Four subunits of nitrate reductase are
encoded by the \emph{narGHJI}genes that are similar to \emph{E. coli}
(Dataset S5) (Moreno-Vivián et al., 1999), and genes involved in Moco
synthesis are not located next to the NAR operon (Fig. 3B).

\subsection*{\texorpdfstring{\emph{Apibacter} lost ancestral gene
families related to pathogenicity and antibiotic
resistance}{Apibacter lost ancestral gene families related to pathogenicity and antibiotic resistance}}

{\label{apibacter-lost-ancestral-gene-families-related-to-pathogenicity-and-antibiotic-resistance}}

Some gene families shared by the common ancestor of \emph{Apibacter} ,
Clade C and E are lost in the \emph{Apibacter} group but are retained in
Clade C and E. As both Clade C and E encompass important mammal and bird
pathogens, the absence of these genes families among
\emph{Apibacter}group suggests that they might be superfluous or
deleterious to the interactions with the host. The 467 gene families
shared by Clade C and E are overrepresented in histidine metabolism,
fatty acid degradation, phenylacetate catabolism and urease activity. A
survey of the genes in these pathways showed that \emph{Apibacter} spp.
lost all genes related to histidine degradation, two key genes involved
in long chain fatty acid beta-oxidation, and the genes responsible for
the production of host toxic virulence using intermediates generated in
the phenylacetate catabolism (Fig. 4) (Teufel et al., 2010). The loss of
genes that are responsible for histidine degradation and long chain
fatty acid beta-oxidation implies that these substrates are less
accessible for\emph{Apibacter} to use as energy sources. To the
contrary, genes of these three pathways are prevalently distributed
among Clade C and E (Dataset S5). All of these pathways are reported to
be involved in host recognization, successful colonization, virulence
factor regulation and production in pathogenic bacteria (Law et al.,
2008; Moraes et al., 2014; Zarzycki-Siek et al., 2013; Zhang, Ritchie,
\& Rainey, 2014).

Antibiotic resistances are promiscuous for bacteria in the
\emph{Flavobacteriaceae} family, causing difficulties in the treatment
of their infection (McBride, 2014). Referring to the CARD database, 13
antibiotic resistance genes which conferred resistance to beta-lactam,
fluoroquinolones, tetracyclines and glycopeptides were identified in
genomes of Clade C and E (Fig S3). These resistance genes are absent in
the \emph{Apibacter} group, except that a lincosamides resistance gene
is identified in \emph{A. adventories} wkB301. These results may be
explained by the fact that\emph{A. cerana} bee gut microbes are less
exposed to antibiotics.

\section*{Discussion}

{\label{discussion}}

Combining FISH and colonization experiments, we revealed the
colonization specificity of \emph{Apibacter} and its distribution in the
bee gut. Comparative genomic analyses of 30 genomes from
the\emph{Flavobacteriaceae} family, including 14 newly
sequenced\emph{Apibacter} genomes from this study and publicly available
genomes for the outgroups, we characterized gene signatures underlying
lifestyle transition and adaptation to bee gut symbionts.

FISH visualization indicates that \emph{Apibacter} coinhabit
with\emph{Snodgrassella} and colonize the epithelium of the bee gut. As
core members of gut bacteria in \emph{A. cerana} , both \emph{Apibacter}
and\emph{Snodgrasella} are microaerophilic, sharing nutritional sources
(Zheng, Powell, Steele, Dietrich, \& Moran, 2017). We showed
that\emph{Apibacter} isolated from \emph{A. cerana} were able to
colonize\emph{A. mellifera} , although at a significantly lower
colonization rate. These results suggest that host incompatibility is
probably not the constraining factor responsible for the rarity of
\emph{Apibacter}in \emph{A. mellifera} . However, it is not yet possible
to examine inter-host competition between \emph{Apibacter} isolated from
different honeybee species, because isolates from \emph{A. mellifera}
were not available to us.

Comparative genomic analysis revealed key gene functions potentially
associated with the adaptation to bee gut niche. In a typical symbiotic
system, benefits provision was considered crucial for the establishment
of a mutualistic relationship (EWALD, 1987; Sachs et al., 2013). Our
findings reveal that \emph{Apibacter} are indeed providing beneficial
traits to the host. For example, genes involved in amino acid
biosynthesis are preserved in \emph{Apibacter} spp., at a background of
overall genome reduction, which echoes those previously reported in
other bee gut symbionts (Kwong et al., 2014). Furthermore,
the\emph{Apibacter} group retained the mannose catabolic gene, which was
responsible for monosaccharide detoxification in the honeybee therefore
broadening food choice for the host (Zheng et al., 2016).

Polysaccharides utilization is a prominent property carried by bee gut
symbionts including \emph{Gilliamella} , \emph{Bifidobacterium}
and\emph{Lactobacillus} (Bonilla-Rosso \& Engel, 2018; Engel, Martinson,
\& Moran, 2012; Ke\selectlanguage{polish}š\selectlanguage{english}nerov\selectlanguage{ngerman}á et al., 2017). However, relevant genes are
substantially lost in the \emph{Apibacter} group. Interestingly, the
core bacterial species \emph{Snodgrasella} that coinhabit
with\emph{Apibacter} at \emph{A. cerana} gut epithelium also lack the
capacity to utilize polysaccharides (Kwong et al., 2014). We speculate
that polysaccharides might be limited in the niche that they share.

The gut lumen is mainly anaerobic, where the dominant symbiotic
anaerobes inhabit. However, oxygen can diffuse from the intestinal
epithelium cells and create a microaerobic environment for facultative
anaerobes (He et al., 1999; Zheng, Powell, et al., 2017). A previous
study found both cytochrome \emph{bd} and \emph{cbb\textsubscript{3}} in
the \emph{Apibacter} genome, which were presumably involved in
microaerobic respiration (Kwong et al., 2018). In the present work, we
identified additional anaerobic respiration NAR operon that was
conserved within the \emph{Apibacter} group and in the
coinhabiting\emph{Snodgrassella} , but absent from the other four core
bee gut bacteria species. These observations suggest that the NAR
pathway might be important for the microbiome to colonize intestinal
epithelium. Such respiratory flexibility might enable \emph{Apibacter}
to survive altered oxygen tensions. This finding is congruent with the
observation in mouse\emph{E. coli} , where they require both
microaerobic and anaerobic respirations for successful colonization
(Jones et al., 2007). A further study proved that the NAR pathway played
a key role in \emph{E. coli}colonization of the mouse gut, because the
NarG mutant showed colonization deficiency for both commensal bacteria
and pathogenic\emph{E. coli} (Jones et al., 2011). These results are in
line with the observation that nitrate reduction could facilitate the
growth of gut microaerobic bacteria at low oxygen conditions (Tiso \&
Schechter, 2015). Therefore, we conclude that the NAR operon is an
important genetic signature for \emph{Apibacter} adaptation to the bee
gut.

Genes that are shared between the LCAs of Clade C and Clade E but absent
from the \emph{Apibacter} group, contain functions either deleterious to
the mutualistic relationship with the host, or redundant for the
symbiotic lifestyle. Histidine biosynthesis is one of the most energy
consuming processes for bacteria, such that the degradation of histidine
as carbon and nitrogen sources is strictly regulated (Bender, 2012). The
histidine catabolism is limited in bee gut environments, as oxygen is
required for the activation of the Hut operon (Goldberg \& Hanau, 1980).
Considering that the bee gut is mostly anoxic, the Hut pathway is highly
likely to be malfunctioning in\emph{Apibacter} and is susceptive to be
lost. In addition, histidine degradation is important for pathogens to
recognize eukaryotic hosts and to activate virulence factors (Zhang et
al., 2014).

In conclusion, combining molecular and colonization experiments, for the
first time, we visualized and quantified the distribution
of\emph{Apibacter} spp. inside the bee gut, and proved
that\emph{Apibacter} isolates of \emph{A. cerana} could survive in
\emph{A. mellifera} . Genomic comparisons with relatives living on other
lifestyles revealed that host beneficial traits and respiration nitrate
reduction (NAR pathway) were key functions for adaptation to the bee gut
environment.

\section*{Acknowledgements}

{\label{acknowledgements}}

The work was funded by the Program of Ministry of Science and Technology
of China (2018FY100403) and National Natural Science Foundation of China
(31772493).

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\section*{Data Accessibility}

{\label{data-accessibility}}

The \emph{Apibacter} genome sequences in this study are available under
NCBI Bioproject accession PRJNA578212.

Supplementary Datasets:

Dataset S1. List of genome sequence deposition and strain collection
sites.

Dataset S2. List of \emph{Apibacter} specific gene families gain and
loss and COG category abbreviations.

Dataset S3. Distribution of CAZy genes according to three groups and
PULs similarities among \emph{Apibacter} group.

Dataset S4. Distribution of amino acid biosynthesis genes distributions.

Dataset S5. List of sublineage specific gene families.

\section*{Author Contributions}

{\label{author-contributions}}

X. Zhang conceived the idea and wrote the manuscript with support from
W. Zhang. W. Zhang performed the experiment and bioinformatic analysis.
X. Zhang and W. Zhang analyzed data. Q. Su and W. Zhang isolated
the\emph{Apibacter} strains used in this work. M. Tang conducted the
assembly of the \emph{Apibacter} genomes. X. Zhou supervised the
findings of this work.

\section*{Figures}

{\label{figures}}

\textbf{Hosted file}

\verb`image1.emf`  available at \url{https://authorea.com/users/322259/articles/451312-genomic-features-underlying-evolutionary-transitions-of-apibacter-to-honeybee-gut-symbionts}

\textbf{Figure 1} Characterization of the\emph{Apibacter} colonization
in honeybees gut. A - D. Localization of\emph{Apibacter} spp.
coordinating with \emph{Snodgrassella} and\emph{Gilliamella} within
midgut and ileum of \emph{A. cerana} worker bees. E. The colonization
abundance of \emph{Apibacter} spp. at different intestinal organs in the
gut of \emph{A. cerana} worker bees. (n=15 per treatment) F.
Colonization of newly emerged germ free bees of\emph{A. cerana} and
\emph{A. mellifera} with \emph{Apibacter} isolated B3706 from \emph{A.
cerana} (n=9). The results of Mann--Whitney U tests (*P \textless{}
0.05) are shown.\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image2/image2}

\end{center}
\end{figure}

\textbf{Figure 2} Genes flux analysis and functional classification of
niche specifying genes. A. Loss and gain of gene families (blue, gain;
red, loss) in gene content since the last common ancestor (numbers in
black). The size of the pie chart reflects the amplitude of total gene
flux (gain + loss). B. COG functional classification of genes lost in
the \emph{Apibacter} genome. See Dataset S2 for complete list of gene
families with annotations and COG category abbreviations. C. Venn
diagram shows gene family distribution among the three major
groups:\emph{Apibacter} group, Clade C and Clade E.\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image3/image3}

\end{center}
\end{figure}

\textbf{Figure 3} Respiratory nitrate reduction pathway is specifically
conserved in the \emph{Apibacter} group, which is also possessed
by\emph{Snodgrassella} . A. Schematic diagram shows that the respiration
nitrate reduction pathway together with molybdate cofactor synthesis are
conserved accoss the \emph{Apibacter} group (pink arrows), and nitrite
detoxification (grey arrows) is absent in the \emph{Apibacter} group but
possessed by genomes in Clade C/E (genes distribution are provided in
Dataset S5). B. Genomic regions of type strains of \emph{Apibacter}
from\emph{A. cerana} , bumble bee and \emph{A. dorsata} encodes genes
involved in respiratory nitrate reduction. Vertical grey blocks connect
homologous genes among type strains, with numbers representing the
percentage of sequence similarities. Genes in pink encodes nitrate
reductase and transporter. Genes in blue encodes molybdate cofactor
synthesis. Genes in grey are hypothetical genes or genes that are not
directly related.

\textbf{Hosted file}

\verb`image4.emf`  available at \url{https://authorea.com/users/322259/articles/451312-genomic-features-underlying-evolutionary-transitions-of-apibacter-to-honeybee-gut-symbionts}

\textbf{Figure 4} Metabolic pathways that are present in genomes of the
Clade C and Clade E, but incomplete among genomes of
\emph{Apibacter}group. A. Histidine degradation pathway; B. Fatty acid
beta-oxidation pathway; C. Phenylacetate oxidation pathway. Black arrow,
genes mostly present in genomes from three groups; Red arrow, genes are
absent among all genomes in \emph{Apibacter} group, but possessed by
genomes in Clade C and E (genes distributions are provided in Dataset
S5). Substrates in red in phenylacetate degradation are documented to be
toxic to host.

\selectlanguage{english}
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