RESEARCH ARTICLE

Discovery and characterization of aprevalent human gut bacterial enzyme
sufficient for the inactivation of a familyof plant toxins
Nitzan Koppel1, Jordan E Bisanz2, Maria-Eirini Pandelia3, Peter J Turnbaugh2,4*,
Emily P Balskus1,5*1

Department of Chemistry and Chemical Biology, Harvard University, Cambridge,
United States; 2Department of Microbiology & Immunology, University of California,
San Francisco, United States; 3Department of Biochemistry, Brandeis University,
Waltham, United States; 4Chan Zuckerberg Biohub, San Francisco, United States;
5
Broad Institute, Cambridge, United States

Abstract Although the human gut microbiome plays a prominent role in xenobiotic*For correspondence:Peter.Turnbaugh@ucsf.edu (PJT);
balskus@chemistry.harvard.edu(EPB)
Competing interest: See
page 26
Funding: See page 26
Received: 29 November 2017
Accepted: 11 April 2018
Published: 15 May 2018
Reviewing editor: Ruth Emily
Ley, Max Planck Institute forDevelopmental Biology,
Germany
Copyright Koppel et al. This
article is distributed under theterms of the Creative Commons
Attribution License, which
permits unrestricted use andredistribution provided that theoriginal author and source arecredited.

transformation, most of the genes and enzymes responsible for this metabolism are unknown.
Recently, we linked the two-gene ‘cardiac glycoside reductase’ (cgr) operon encoded by the gut
Actinobacterium Eggerthella lenta to inactivation of the cardiac medication and plant natural
product digoxin. Here, we compared the genomes of 25 E. lenta strains and close relatives,
revealing an expanded 8-gene cgr-associated gene cluster present in all digoxin metabolizers andabsent in non-metabolizers. Using heterologous expression and in vitro biochemical
characterization, we discovered that a single flavin- and [4Fe-4S] cluster-dependent reductase,Cgr2, is sufficient for digoxin inactivation. Unexpectedly, Cgr2 displayed strict specificity for
digoxin and other cardenolides. Quantification of cgr2 in gut microbiomes revealed that this geneis widespread and conserved in the human population. Together, these results demonstrate thathuman-associated gut bacteria maintain specialized enzymes that protect against ingested plant
toxins.
DOI: https://doi.org/10.7554/eLife.33953.001

Introduction
The human gut microbiome extends the metabolic capabilities of the human body, extracting
energy from otherwise indigestible dietary polysaccharides, synthesizing essential vitamins andamino acids, and modifying endogenous compounds. This microbial community also extensively
metabolizes xenobiotics, including synthetic and natural product-based drugs, food additives, andenvironmental toxins (Koppel et al., 2017; Spanogiannopoulos et al., 2016). Previous work has
focused on cataloguing the xenobiotics subject to transformation by gut microbes, their downstream
products, and the extensive inter-individual variation in these activities (Koppel et al., 2017;
Spanogiannopoulos et al., 2016). However, reduction of these complex metabolic networks tomechanism has been limited by our lack of knowledge about the microbial enzymes responsible forxenobiotic biotransformation.
Digoxin, a cardenolide used to treat heart failure and arrhythmia, represents a valuable test casefor our ability to elucidate the biochemical and evolutionary underpinnings of gut microbial xenobiotic metabolism. It has been known for decades that human gut bacteria reduce digoxin to the inactive metabolite dihydrodigoxin, decreasing drug efficacy and toxicity (Saha et al., 1983;

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eLife digest Trillions of microbes live within the human gut and influence our health. In
particular, these microbes can modify food and drugs into compounds (metabolites) that humans
cannot produce on their own. These compounds are often beneficial to the human host, but in somecases – for example, if the modification alters how a drug works – can be detrimental.
Digoxin is a toxic chemical produced by plants that, in low doses, can be used to treat heart
conditions. It has been known for decades that the human gut bacterium Eggerthella lenta
transforms digoxin into a metabolite that is an ineffective drug. Microbes use biological catalysts
called enzymes to produce metabolites, but it was not known which enzymes enable E. lenta tomodify digoxin.
Using biochemical and genomic techniques, Koppel et al. now show that an enzyme called Cgr2
inactivates digoxin and other related plant toxins. Data about the gut microbes in nearly 1,900
people from three continents revealed that bacteria that can produce Cgr2 were present in the gutsof more than 40% of the individuals, although often in low abundance. Further experiments did notreveal any obvious benefits that E. lenta gains from modifying digoxin. Instead, Koppel et al.
propose that the bacteria carry out this modification to protect their human host from plant toxins.
The results presented by Koppel et al. emphasise that the activities of gut microbes should beconsidered when designing new drugs or assessing how they work in the human body. The
strategies used to identify Cgr2 could now be applied to discover other important gut microbe-druginteractions. Ultimately, this knowledge will help us to predict and control the activities of gut
microbes in ways that could improve human health.
DOI: https://doi.org/10.7554/eLife.33953.002

Lindenbaum et al., 1981a; Lindenbaum et al., 1981b). Screening hundreds of gut bacterial strainsfrom humans that excreted high levels of dihydrodigoxin revealed only two isolates that were capable of metabolizing digoxin, both of which were strains of the anaerobic, low abundance bacterium
Eggerthella lenta (Saha et al., 1983). However, the presence of E. lenta in the gut microbiome cannot accurately predict this reactivity, as patients colonized by E. lenta still show marked variation in
dihydrodigoxin production (Mathan et al., 1989; Alam et al., 1988). We identified two mechanisms
underlying this discrepancy: strain-level variations in the E. lenta population and inhibition of bacterial drug metabolism by dietary amino acids. Digoxin induces expression of a 2-gene operon
encoded by the type strain of E. lenta (DSM 2243), which we named the cardiac glycoside reductase(cgr) operon (Haiser et al., 2013). The cgr operon was absent in two E. lenta strains that did notmetabolize digoxin (‘non-reducing’ strains) and cgr operon presence and abundance predicted theextent of drug inactivation by human gut microbial communities in ex vivo incubations (Haiser et al.,2013). Furthermore, germ-free mice that had been mono-colonized by a reducing (cgr+) strain of E.
lenta had lower serum levels of digoxin than mice colonized by a non-reducing (cgr-) strain, and dietary arginine efficiently blocked digoxin reduction by the cgr operon in cgr+ E. lenta-colonized mice
(Haiser et al., 2013; Haiser et al., 2014).
Although it is tempting to consider applying these insights to develop novel microbiome-based
diagnostics and co-therapies (Spanogiannopoulos et al., 2016), multiple critical questions remained
unaddressed. Our original studies were entirely based on the E. lenta type strain, isolated in 1938
from a rectal cancer biopsy (Moore et al., 1971); thus, the presence of cgr +E. lenta in the modernday human gastrointestinal tract was unclear. Although we had associated the cgr operon withdigoxin reduction, the minimal genetic machinery necessary and sufficient for this biotransformation
had not been determined. Perhaps most importantly, the specificity of the digoxin-reducing enzyme(s) for cardenolides and their ability to accept additional endogenous or ingested substrates
remained unclear.
Here, we used a combination of comparative genomics, heterologous expression, biochemistry,
and metagenomics to address these long-standing questions. We uncovered a highly conserved
cluster of genes that co-occurs with the cgr operon, representing a single genetic locus predictive of
digoxin metabolism. We demonstrated that a single protein encoded by this locus, Cgr2, is sufficientfor digoxin reduction and is widespread in human gut microbiomes. Cgr2 is a novel oxygen-sensitive

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*

*

*

*

*
*
*

cgr2
cac1
cac3
cac4
cac5
cac6
Elen_0356
Elen_0640
Elen_0639
hypothetical
Elen_0281
Elen_2443

Relative base position (kb)
5
10
15

0
El1
El2
El3
El4
El5
El6
El7
El8
El9
El10
El11
El12
El13
El15
El16
El17
El18
El19
El20
El21

20

100
95
90
85
80

cgr-associated gene cluster

E

cgr1

Elen_2441*

Elen_2440

2957500

cgr2 cac1 cac2 cac3

cac4

cac5 cac6*
2960000 2962500 2965000
DSM 2243 base position

Strains
Gene present Gene absent

2967500
7.5
0.0

log2(FC)

100
50
0% Digoxin reduction

Non-red. Ortholog D
clustercgr1

Global nucleotide
identity to DSM 2243*

C

Reducer
El17 86
100
El10
1
100
El2
62
El1
El13
El19 63
El14 97
100
El3
100
El7
5
El15 92
96
100
El8
50
El12
100
El5
100
El6
El9
100
El18
93
El16
100
100
10
El4
El11
100
El20
100
El21
100
Es1
100
Ph1
Gs1
Gs2
MeOH
Non-reducer
15
BHI
Reducer

E. lenta strain

B

Ranked ImportanceA

Microbiology and Infectious Disease

Figure 1. Comparative genomics expands the boundaries of the cgr operon. (A) Survey of digoxin reduction in 21 strains of E. lenta (El#), 2 strains ofGordonibacter spp. (Gs#), E. sinensis (Es1), and Paraeggerthella hongkongesis (Ph1) (Figure 1—source data 1 and 2) revealed eight strains capable ofreducing digoxin to dihydrodigoxin (*p<0.05, ANOVA with Dunnett’s test vs. vehicle controls). Data represents mean ± standard error of the mean
(SEM) over three biological replicates. (B) Digoxin reduction did not correlate with phylogeny in E. lenta species (cladogram displayed with bootstrap
values indicated at nodes; p=0.275, K = 0.049, Blomberg’s K). (C) Comparative genomics using a random forest classifier (see Materials and methods)
revealed seven genes with perfect predictive accuracy for digoxin reduction. The orthologous cluster identified as hypothetical corresponds to an open
reading frame present at position 299442..2995131 in the DSM 2243 reference genome. (D) Analysis of genomic context revealed a highly conserved
10.4 kb locus of 7 genes that flank a short, conserved hypothetical gene, herein termed the cgr-associated gene cluster (cac). (E) Analysis of gene
expression in the cgr-associated gene cluster revealed only the cgr-locus was significantly upregulated by exposure to digoxin. * FDR < 0.1 (Figure 1—
source data 3).
DOI: https://doi.org/10.7554/eLife.33953.003
The following source data is available for figure 1:
Source data 1. Bacterial strains used in study.
DOI: https://doi.org/10.7554/eLife.33953.004
Source data 2. Digoxin Reduction by E.lenta and related bacterial isolates.
DOI: https://doi.org/10.7554/eLife.33953.005
Source data 3. Differential gene expression analysis of cgr-associated genes in E.lenta DSM 2243.
DOI: https://doi.org/10.7554/eLife.33953.006

reductase that requires flavin adenine dinucleotide (FAD) and at least one [4Fe-4S] cluster. Surprisingly, we found that Cgr2 only accepts digoxin and other cardenolides, prompting the provocative
hypothesis that the gut microbiome provides a first-line of protection against ingested toxins analogous to that of host enzymes expressed in the intestinal epithelium and liver. Finally, this work establishes a generalizable framework for mechanistic investigations of gut microbial xenobiotic
metabolism that will enhance our understanding of the complex dietary, host, and microbial factorsthat impact pharmacology and toxicology, and provide a stronger foundation for translational studies in patient populations.

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Results
Identification of a single genetic locus conserved in all digoxin-reducing
strains
E. lenta strains vary in their ability to reduce digoxin (Haiser et al., 2013; Haiser et al., 2014); however, our prior attempts at identifying the minimal genetic machinery necessary for metabolism werelimited by the availability of just a single strain capable of this activity (E. lenta DSM 2243). Throughpublic repositories and isolation of novel strains, we curated, sequenced, and annotated a collectionof 25 E. lenta and closely related Coriobacteriia strains (Bisanz et al., 2018). These bacteria wereisolated from 22 individuals in 6 countries across three continents spanning the years of 1938–2015(Figure 1—source data 1).
We used liquid chromatography-tandem mass spectrometry (LC-MS/MS) to quantify the biotransformation of digoxin to dihydrodigoxin by each strain and identified seven additional strains capableof drug inactivation (Figure 1A). Culturing experiments were performed using 10 mM of digoxin,
which is within the estimated range (0.4–14.6 mM) of therapeutic concentrations of the drug in thegut prior to absorption by host epithelial cells (Schiller et al., 2005). The digoxin metabolizingstrains did not exhibit a significant phylogenetic signal (Figure 1B; p=0.275, K = 0.049, Blomberg’s
K test), suggesting that this phenotypic trait has been gained (or lost) multiple times over the courseof E. lenta evolution. Machine learning (random forests) analysis of orthologous gene cluster presence/absence across the strain collection revealed a single genetic locus with 100% discriminativevalue between metabolizers and non-metabolizers (Figure 1C). This locus, referred to hereafter as
the cgr gene cluster, includes the previously identified 2-gene cgr operon (cgr1 and cgr2) and sixneighboring genes termed cac (cgr-associated cluster) genes (Figure 1D). The cac genes include aputative LuxR type transcriptional regulator (Cac3), a predicted flavin-dependent fumarate reductase(Cac4), three proteins of unknown function (Cac1, Cac5, Cac6), and a short protein (Cac2) that isconserved in both cgr- and cgr+ strains of E. lenta.
The 10.4 kb cgr gene cluster was highly conserved between strains with an average global nucleotide identity of 99.95 ± 0.05% (mean ± standard deviation (SD)). Both cgr+ and cgr- strains share a
short 174 bp hypothetical gene (cac2) that is conserved with 100% global nucleotide identity in cgr+ strains, while cgr- strains are 90.20–91.37% identical to cgr+ cac2. This conservation and genomic
context may be indicative of multiple translocations in the region creating the cgr-associated genecluster although obvious markers of recent translocation of the cgr gene cluster are absent.

Cgr2 is sufficient for digoxin reduction
Multiple lines of evidence suggested that the cgr operon encodes the enzymes responsible fordigoxin metabolism. Of the eight genes in the cgr gene cluster, only three show primary sequencehomology to reductases: cgr1, cgr2, and cac4. RNA sequencing (Haiser et al., 2013) demonstrated
that the cgr operon (cgr1 and cgr2) is highly upregulated (>165 fold) in response to digoxin, whereas
cac4 is not significantly induced (1.3-fold change relative to vehicle controls, p=0.83) (Figure 1E).
The remainder of the cgr-associated cluster is largely transcriptionally dormant during exponential
growth both with and without the presence of digoxin (<6 normalized counts per gene) andis therefore unlikely to be linked to digoxin metabolism. Initial annotations of Cgr1 and Cgr2 suggested both proteins might mediate digoxin reduction. Cgr1 is a putative membrane-anchored protein that belongs to the cytochrome c3 superfamily (Pfam 14537) and is predicted to harbor
covalently bound heme groups (CXXCH motif). It most closely resembles the NapC/NirT (NrfH) family of proteins that transfer electrons from the membrane quinone pool to associated reductases,
facilitating reduction of terminal electron acceptors such as nitrite and sulfite (Kemp et al., 2010;
Kern et al., 2008). We also identified a close homolog of Cgr1 (Elen_2528) in E. lenta DSM 2243
(91.75% amino acid identity, BLASTP) that is a component of the E. lenta core genome (99.39 ±
0.81% global identity mean ± SD). The presence of this highly similar protein in both metabolizing
and non-metabolizing strains further indicates that Cgr1 is involved in a more general function (e.g.
electron transfer, membrane docking) rather than direct reduction of digoxin. On the other hand,Cgr2 is unique to the genomes of cgr+ E. lenta, and the closest homologs of Cgr2 display <28%amino acid identity. Cgr2 is a homolog of flavin adenine dinucleotide (FAD)-dependent fumarate
reductases (Pfam 00890; Interpro 003953/027477) and is predicted to undergo secretion via the twin

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Figure 2. Cgr2 is sufficient for digoxin reduction and requires FAD and [4Fe-4S] cluster(s) for activity. (A) Whole cell assays using R. erythropolis
expressing Cgr1 and Cgr2 constructs demonstrated that Cgr2 is sufficient for reducing digoxin. Data represents the mean ± SEM (n = 3 biologicalreplicates). Asterisks indicate statistical significance of each variant as compared to empty vector by Student’s t test (**p<0.01, ***p<0.001) (Figure 2—
source data 1). (B) Annotation and amino acid numbering of Cgr2, including the predicted Tat secretion signal and three conserved flavin-binding
motifs from the glutathione reductase family (X = any amino acid; h = hydrophobic residue). (C) In vitro activity of Cgr2 for digoxin reduction using
reduced methyl viologen as an electron donor, analyzed by liquid chromatography-tandem mass spectrometry (LC-MS/MS). [Fe-S] cluster
reconstitution, FAD, and anaerobic conditions are required for Cgr2 activity. Data represents the mean ± SEM (n = 3 independent experiments)(Figure 2—source data 2). FAD = flavin adenine dinucleotide; FMN = flavin mononucleotide. (D) Ultraviolet-visible (UV-Vis) absorption spectra of Cgr2
revealed an oxygen-sensitive peak centered around 400 nm that increased upon [Fe-S] cluster reconstitution, supporting the presence of [4Fe-4S]
clusters in Cgr2. (E) Electron paramagnetic resonance (EPR) spectra of sodium dithionite-reduced Cgr2 reconstituted with iron ammonium sulfate
hexahydrate ((NH4)2Fe(SO4)26H20) and sodium sulfide (Na2S9H20). G-values and decreased EPR signal intensity at higher temperatures (10 – 40 K)
indicated the presence of low potential [4Fe-4S]1+ clusters. Experimental conditions were microwave frequency 9.38 GHz, microwave power 0.2 mW,
modulation amplitude 0.6 mT, and receiver gain 40 dB.
DOI: https://doi.org/10.7554/eLife.33953.007
The following source data and figure supplements are available for figure 2:
Source data 1. Digoxin metabolism by R. erythropolis overexpressing Cgr proteins.
DOI: https://doi.org/10.7554/eLife.33953.011
Source data 2. Digoxin metabolism by Cgr2 in vitro.
DOI: https://doi.org/10.7554/eLife.33953.012
Source data 3. Examples of [2Fe-2S], [3Fe-4S], and [4Fe-4S] cluster binding motifs that are not found in Cgr2 (Zhang et al., 2010; NakamaruOgiso et al., 2002; Lee et al., 2004; Pandelia et al., 2011; Schnackerz et al., 2004; Leech et al., 2003; Gorodetsky et al., 2008; Lee et al., 2010;
Weiner et al., 2007; Klinge et al., 2007; Dickert et al., 2002; Conover et al., 1990; Schneider and Schmidt, 2005; Iwasaki et al., 2000;
Banci et al., 2013; Dailey and Dailey, 2002; Jung et al., 2000).
DOI: https://doi.org/10.7554/eLife.33953.013
Source data 4. Digoxin metabolism by R. erythropolis overexpressing Cgr2 cysteine to alanine point mutants.
DOI: https://doi.org/10.7554/eLife.33953.014
Source data 5. Digoxin metabolism by Cgr2 cysteine to alanine point mutants in vitro.
DOI: https://doi.org/10.7554/eLife.33953.015
Figure supplement 1. [Fe-S] cluster(s) affect Cgr2 stability and oligomerization.
DOI: https://doi.org/10.7554/eLife.33953.008
Figure supplement 2. Cgr2 activity, but not EPR-active [4Fe-4S] clusters, increase with higher Fe and S equivalents.
DOI: https://doi.org/10.7554/eLife.33953.009
Figure supplement 3. Identification of 6 cysteine residues important for Cgr2 activity.
Figure 2 continued on next page

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Figure 2 continued
DOI: https://doi.org/10.7554/eLife.33953.010

arginine translocation (Tat) pathway. Taken together, these observations led us to hypothesize that
Cgr1 and Cgr2 form a membrane-associated complex that catalyzes reduction of the a,b-unsaturated butyrolactone of digoxin.
Heterologous expression of Cgr1 and Cgr2 in the model Actinobacterium Rhodococcus erythropolis L88 (Mitani et al., 2005; Nakashima and Tamura, 2004a, 2004b) allowed us to test whether
these proteins were sufficient for digoxin reduction. After inducing protein expression, cultures wereincubated with 10 mM of digoxin and dihydrodigoxin production was quantified by LC-MS/MS(Figure 2A). The Cgr2 expressing strains showed a significant increase in dihydrodigoxin levels relative to empty vector controls (Figure 2A; Figure 2—source data 1). In contrast, no activity wasobserved for the strain expressing only Cgr1, although this could be due to a lack of protein, as nooverexpression of Cgr1 could be detected in either clarified lysates or membrane fractions. These
results show that Cgr2 is sufficient for digoxin reduction in R. erythropolis cells, and endogenous
redox active proteins and/or metabolites in this heterologous host may fulfill the putative function ofCgr1 as an electron donor.

Cgr2 activity depends on [4Fe-4S] cluster(s)Having identified Cgr2 as the critical reductase enzyme, we next aimed to reconstitute its activity invitro. Examining multiple tagged versions and truncations of Cgr2 in R. erythropolis revealed that a
Cgr2(–48aa)-NHis6-tagged construct lacking the Tat secretion signal gave the highest yield andpurity (Figure 2—figure supplement 1A). This construct, hereafter referred to as ‘wild-type’ Cgr2,
was used for all in vitro studies. Computational analysis of Cgr2 predicted that it would bind flavinthrough a Rossmann fold (Dym and Eisenberg, 2001), and the three motifs required for cofactor
binding are conserved in all 8 Cgr2 sequences (Figure 2B). However, Cgr2 did not co-purify with flavin. Moreover, Cgr2 obtained from initial purifications was thermally unstable (melting
temperature <37˚C), was prone to degradation during cell lysis (Figure 2—figure supplement 1),
and displayed low activity for digoxin reduction (Figure 2C); Figure 2—source data 2). Together,
these observations indicated that an essential cofactor was likely missing. We also noticed that purified Cgr2 was light brown in color, suggesting the presence of a metallocofactor. Certain flavindependent reductases use metallocofactors to mediate the transfer of electrons to the active site,
including cytochromes c in soluble enzymes and oxygen-sensitive iron-sulfur ([Fe-S]) clusters in membrane-bound enzymes (Kern et al., 2008; Iverson et al., 2002; Leys et al., 1999). The combinationof the brown color and the presence of 16 cysteines in the mature Cgr2 sequence led us to hypothesize that this enzyme contained one or more [Fe-S] clusters. However, we were unable to detect any
canonical [2Fe-2S], [3Fe-4Fe], or [4Fe-4S] cluster binding motifs within the Cgr2 sequence (Figure 2—
source data 3) (Zhang et al., 2010; Nakamaru-Ogiso et al., 2002; Lee et al., 2004;
Pandelia et al., 2011; Schnackerz et al., 2004; Leech et al., 2003; Gorodetsky et al., 2008;
Lee et al., 2010; Weiner et al., 2007; Klinge et al., 2007; Dickert et al., 2002; Conover et al.,
1990; Schneider and Schmidt, 2005; Iwasaki et al., 2000; Banci et al., 2013; Dailey and Dailey,2002; Jung et al., 2000).
We therefore sought to determine whether Cgr2 required an [Fe-S] cluster to catalyze digoxin
reduction. Attempts to chemically reconstitute [Fe-S] cluster formation by incubating Cgr2 with ironand sulfide under anaerobic conditions greatly improved protein stability (Figure 2—figure supplement 1B–D). In addition to performing this reconstitution step, including FAD in assay mixtures dramatically increased the digoxin reduction activity of purified protein (Figure 2C). Even afterreconstitution, Cgr2 required excess FAD for maximal activity, suggesting that Cgr1 could be important for enhancing FAD binding, as has been observed for proteins whose FAD-binding site is predicted to occur at the interface of two domains (Kleven et al., 2015).
Having demonstrated that [Fe-S] reconstitution was essential for activity, we next attempted todetermine the exact nature of this metallocofactor. Prior to reconstitution, purified Cgr2 contained
between 0.2–0.6 equivalents of iron and sulfide, and its ultraviolet-visible (UV-vis) spectrum displayed absorption features consistent with the presence of low levels of [Fe-S] clusters (Ayala-

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Castro et al., 2008) (Figure 2D). After reconstitution, Cgr2 exhibited a broad peak around 400 nm(Figure 2D) that decreased in absorbance upon addition of an excess amount of the reducing agent
sodium dithionite (Figure 2—figure supplement 2A). These spectral properties are characteristic ofredox-active [Fe-S] clusters. Exposure of reconstituted Cgr2 to oxygen led to [Fe-S] cluster decomposition as evidenced by a decrease in the absorbance at 400 nm (Figure 2D), demonstrating thatthe [Fe-S] cluster(s) of Cgr2 are oxygen-sensitive.
Although these UV-vis experiments indicated the presence of [Fe-S] cluster(s) in Cgr2, they could
not define the precise structures of these cofactors. To more definitively characterize these metallocofactors, we turned to electron paramagnetic resonance (EPR) spectroscopy. This technique detects
unpaired electrons and can differentiate between the various types of [Fe-S] clusters as well as provide information about cluster orientation and redox state. In the absence of a reducing agent, purified, unreconstituted Cgr2 does not have an EPR signal (Figure 2—figure supplement 2B). Upon
reduction with sodium dithionite, the EPR spectrum of Cgr2 exhibits a signal with axial symmetry
and with principal g-components of 2.045 and 1.94. This signal increases in intensity upon reconstitution of Cgr2 (10 K), and is barely detectable above 40 K (Figure 2E). Both the principal g-values andthe relaxation properties (temperature dependence) of this signal are characteristic of low-potential
tetranuclear [4Fe-4S]1+ centers. These observations indicate that Cgr2 contains [4Fe-4S]2+ clusterthat can undergo reduction to the corresponding [4Fe-4S]1+ state. These redox properties suggestthat these [4Fe-4S] cluster(s) may participate in catalysis (electron transfer).
Next, we sought to determine how many [4Fe-4S] clusters were present in Cgr2. Using a Cu2+EDTA standard, we determined that purified, unreconstituted Cgr2 contains 0.02–0.03 [4Fe-4S]1+clusters per protein monomer. After reconstitution with iron and sulfide, the intensity of the EPR signal increased to 0.13–0.25 [4Fe-4S]1+ clusters per Cgr2. Though these data may suggest the presence of one [4Fe-4S] cofactor per Cgr2, they do not exclude the possibility of multiple [Fe-S]
centers. Indeed, the in vitro activity of Cgr2 increases upon reconstitution with increasing equivalentsof iron and sulfide (Figure 2—figure supplement 2C–D). Additional spectroscopic or structural characterization (e.g. crystallography) will be required to definitively determine the number of [Fe-S] cluster(s) present in Cgr2.

Identification of amino acids required for Cgr2 function
As motif analysis could not identify putative [4Fe-4S] cluster binding sites in Cgr2, we attempted touse site-directed mutagenesis to reveal the cysteine residues required for cofactor assembly. Individually mutating each of the 16 cysteines present in wild-type Cgr2 to alanine revealed six residues
that, when mutated, significantly decreased dihydrodigoxin production by both heterologously
expressed and purified Cgr2 (Figure 2—figure supplement 1A; Figure 2—figure supplement 3A–
C; Figure 2—source data 4; Figure 2—source data 5). EPR analysis of these six Cgr2 mutants
revealed comparable levels of [4Fe-4S]1+ cluster incorporation relative to the wild-type enzyme (Figure 2—figure supplement 3D), which may argue against the involvement of these cysteines in [4Fe4S] cluster ligation. However, substitution of a single cysteine residue may not always be sufficient toprevent [4Fe-4S] cluster formation (Iismaa et al., 1991; Hewitson et al., 2002; Martı́n et al., 1990).
Alternatively, these six cysteines may be critical for protein structure (e.g. through participating in
disulfide formation) or could coordinate another metal center not detectable in our spectroscopic
experiments. Consistent with this latter proposal, we found that a range of divalent metal cations(Fe2+, Mn2+, Mg2+) stimulated the activity of Cgr2 in vitro (Figure 2—figure supplement 3E) without
altering protein stability. Additionally, Fe2+ stimulated the in vitro activity of only 3 out of 6 impaired
mutants (C158A, C187A, C327A) (Figure 2—figure supplement 3F). Notably, binding of digoxin to
its target in human cells, Na+/K+ ATPase, is thought to be mediated by long-range electrostaticinteractions between a Mg2+ ion and the electron rich, partially negatively charged oxygen atom of
the unsaturated lactone (Laursen et al., 2015; Weigand et al., 2014). It is possible that the threeremaining cysteine residues (C82, C265, C535) could influence binding of a divalent metal cationthat similarly positions or activates digoxin in the Cgr2 active site.
To identify additional amino acids that may be important for Cgr2 function, we compared theCgr2 sequences encoded within our collection of E. lenta genomes. Strikingly, only two cgr2 nucleotide variants were detected, which were validated by targeted Sanger sequencing. One of these variants is only found in the DSM 2243 type strain resulting in a conservative methionine (M) to valine(V) substitution at position 381. The other results in either aromatic tyrosine (Y) as in the type strain

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Figure 3. A single polymorphism in Cgr2 at position 333 (Y/N) leads to altered metabolism of digoxin. (A) Analysisof Cgr2 amino acid sequence composition of isolate genomes (n = 8) and reconstructed sequences from gut
microbiome datasets (n = 14) revealed a single non-conservative Y333N variant in isolate strains supported bymetagenomes (12Y/10N). (B) Nucleotide variation in the cgr-associated gene cluster. Reads aligned from bothisolate genomes (iso) and metagenomes were aligned to the DSM 2243 reference assembly and plotted if there
was coverage of the Y333N variant position (CP001726.1: 2959294 bp). Variants were called when at least one read
was mapped to the position and > 50% of reads supported an alternative base. Read depth at any given position
is indicated by shading. Confirming assembly-based methods, cgr2 amino acid position 333 was bi-allelic with 5 of
8 isolate genomes and 15 of 49 metagenomes showing the N333 variant (four metagenomes have evidence ofboth alleles) and minimal variation in other regions of the cluster. (C) Average amino acid conservation in the E.
lenta core (n = 1832) and non-singleton accessory genome (n = 2557) demonstrates that cgr2 is in the 67th
percentile for conservation in the pan-genome (78.8th in the core genome, and 58.5th in the non-singleton
accessory genome) with higher average conservation observed in the core genome (98.6 ± 2.41% core, 97.4 ± 5.6%
accessory, mean ± SD). (D) Comparison of digoxin metabolism in culture by E. lenta cgr2- (n = 13 strains),
Cgr2Y333 (n = 3 strains), and Cgr2N333 (n = 5 strains). Control refers to digoxin in BHI media. Each point
represents the mean percent conversion to dihydrodigoxin of each individual strain cultured in biologicaltriplicate. Bars represent the mean ± SEM percent conversion per E. lenta group. Statistical significance between
Y333 and N333 groups was calculated using two-tailed Welch’s t test (p=0.052) (Figure 3—source data 1). (E)Michaelis–Menten kinetics of Cgr2 towards digoxin revealed that the Y333 variant is significantly more active thanthe N333 variant. Data represents mean ± SEM (n = 3 independent experiments) (Figure 3—source data 2;
Figure 3—source data 4). (F) In vitro time course (0 – 4.5 hr) of the conversion of digoxin to dihydrodigoxin by
Cgr2 Y333 and N333 variants. Reaction aliquots were quenched in methanol and analyzed by liquid
chromatography-tandem mass spectrometry. Values represent mean ± SEM (n = 3 independent experiments).
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Figure 3 continued
Asterisks indicate statistical significance at each timepoint of Y333 vs. N333 percent conversion, by Student’s t test(*p<0.05, ***p<0.001) (Figure 3—source data 3).
DOI: https://doi.org/10.7554/eLife.33953.016
The following source data and figure supplement are available for figure 3:
Source data 1. Whole cell activity of E. lenta strains with Y333 vs N333 Cgr2 variants.
DOI: https://doi.org/10.7554/eLife.33953.017
Source data 2. Kinetics of Y333 and N333 Cgr2 variants towards digoxin.
DOI: https://doi.org/10.7554/eLife.33953.019
Source data 3. In vitro time course of digoxin reduction by Y333 and N333 Cgr2 variants.
DOI: https://doi.org/10.7554/eLife.33953.020
Source data 4. Comparison of Cgr2 kinetics with related enzymes towards their respective substrates(Kemp et al., 2010; Rohman et al., 2013; Morris et al., 1994; Bogachev et al., 2012).
DOI: https://doi.org/10.7554/eLife.33953.021
Figure supplement 1. Phylogenetic tree of assayed E. lenta strains showing cgr2 Y333/N333 variants.
DOI: https://doi.org/10.7554/eLife.33953.018

DSM 2243 or neutral asparagine (N) at position 333 (Figure 3A). We were also able to fully or partially reconstruct 14 additional cgr2 sequences using reads mapping to the cgr gene cluster from 96
gut microbiome datasets with a high abundance of E. lenta (>1x coverage or >0.001 proportional
abundance). These metagenome fragments confirmed the presence of both Y333 and N333 variants
in a 9:5 ratio (Figure 3A) while the DSM 2243 M381 variant was not observed. To avoid biases
against lower E. lenta coverage metagenomes, we also applied an assembly-free method based on
calling variants from aligned reads (Figure 3B). This uncovered 49 metagenomes with at least one
read mapping over the variant position, confirming the bi-allelic nature with 15 Y333 and 34 N333
metagenomes. Nearly all metagenomes (41/42) with reads mapping to position 381 supported thevaline residue suggesting that the DSM 2243 M381 variant is rare. Given that this analysis confirmed
the highly conserved nature of the cgr locus, we analyzed the conservation of cgr2 in the context of
the E. lenta pan-genome (based on 24 sequenced isolates) finding that it is at the 67th percentile ofconservation. These results suggest that cgr2 sequence conservation is not unusual for this species,
with the caveat that relatively few genomes were available for analysis (Figure 3C).
Given the ubiquity of variation at position 333, we assessed its functional consequences by comparing the activity of the two Cgr2 variants in vivo and in vitro. E. lenta strains encoding the N333
variant show a trend towards a decreased ability to metabolize digoxin as compared to Y333-encoding strains (Figure 3D; Figure 3—source data 1; p=0.052 Welch’s t-test). This decreased activity
was more readily apparent following incubation of digoxin with Cgr2 proteins in vitro(Figure 3E; Figure 3—source data 2). While kinetic parameters for wild-type Cgr2 (Y333) were
KM = 94.6 ± 7.1 mM and a catalytic efficiency of 2.4 ± 0.8103 M 1 s 1, saturating Vmax conditions
could not be reached for the N333 variant within the range of concentrations where digoxin is soluble (0.5 mM). Despite its lower activity, Cgr2 N333 converted digoxin to dihydrodigoxin at a comparable efficiency to Y333 after 4.5 hr (Figure 3F; Figure 3—source data 3). Compared to theactivity of other FAD-dependent reductases towards their native substrates, the Y333 Cgr2 variant isless efficient for digoxin reduction (Kemp et al., 2010; Rohman et al., 2013; Morris et al., 1994;
Bogachev et al., 2012) (Figure 3—source data 4). This decreased activity could arise from impaired
cofactor binding or reconstitution, or inefficient electron transfer in vitro in the absence of Cgr1.
Alternatively, these results could indicate that digoxin is not the endogenous substrate of Cgr2.

Cgr2 is a novel enzyme that preferentially reduces cardenolides
To systematically test for additional Cgr2 substrates, we assessed the enzyme’s activity toward 28
small molecules using a colorimetric assay (Figure 4; Figure 4—figure supplement 1). These molecules were selected based on their chemical similarity to digoxin and their relevance in the context
of the human gut. Cgr2 displayed robust activity only toward cardenolides, the family of plant toxinsthat includes the pharmaceutical agents digoxin and digitoxin as well as ouabain, which is used as anarrowhead poison (Michalak et al., 2017). The cardenolide aglycones digoxigenin and ouabagenin
were metabolized at a significantly faster rate than their glycosylated forms digoxin and ouabain,

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Figure 4. The substrate scope of Cgr2 is restricted to cardenolides. Rate of methyl viologen oxidation coupled to substrate reduction by Cgr2. Colors
denote different substrate classes. With the exception of the cardenolides, a representative substrate structure is shown. Values represent mean ± SEM(n = 3 independent experiments). **p<0.01, Student’s t test (Figure 4—source data 1). The heatmap generated in ChemMine (Backman et al., 2011)
represents the structural similarity of each compound relative to digoxin. Structural distance matrix is calculated as (1- Tanimoto coefficient), where
lower values represent more structurally similar compounds.
DOI: https://doi.org/10.7554/eLife.33953.022
The following source data and figure supplements are available for figure 4:
Source data 1. Rate of methyl viologen oxidation coupled to substrate reduction by Cgr2.
DOI: https://doi.org/10.7554/eLife.33953.025
Figure supplement 1. Putative substrates for Cgr2 in the context of the human gut.
DOI: https://doi.org/10.7554/eLife.33953.023
Figure supplement 2. Digoxin and related cardenolides do not affect E. lenta growth in rich or minimal media.
DOI: https://doi.org/10.7554/eLife.33953.024

respectively (**p<0.01, Student’s t test). The isolated lactone 2(5H)-furanone was minimally processed by Cgr2, indicating that an intact steroid core is important for activity. However, the qualitatively similar rates observed for reduction of the various cardenolides demonstrates that the enzyme
tolerates differences in the number and position of hydroxyl groups on the steroid scaffold. Overall,

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Figure 5. Sequence similarity network (SSN) analysis reveals that the gut bacterial enzyme Cgr2 is a highly distinct member of a large enzyme family
that is widespread in gut microbes. The SSN was constructed using the top 5000 most similar proteins to Cgr2 from the UniprotKB database. Nodes
represent proteins with 100% sequence identity. (A) SSN displayed with an e-value threshold of 10 50. The seven previously characterized enzymes(PDB ID: 1D4D, 1E39; UniProtKB ID: Q07WU7, Q9Z4P0, 8CVD0, P71864, Q7D5C1) and Cgr2 are colored according to biochemical function. (B) SSN
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Figure 5 continued
displayed with an e-value threshold of 10 130. All nodes that co-clustered with characterized enzymes are shown in the same color, denoting putative
isofunctional activity. With the exception of Cgr2, if a node comes from a gut bacterium, it is colored red rather than the color of the corresponding
cluster.
DOI: https://doi.org/10.7554/eLife.33953.026
The following source data and figure supplements are available for figure 5:
Source data 1. Characterized enzymes within the Cgr2 sequence similarity network.
DOI: https://doi.org/10.7554/eLife.33953.031
Figure supplement 1. Multiple sequence alignment of fumarate reductases.
DOI: https://doi.org/10.7554/eLife.33953.027
Figure supplement 2. Multiple sequence alignment of urocanate reductases.
DOI: https://doi.org/10.7554/eLife.33953.028
Figure supplement 3. Multiple sequence alignment of ketosteroid dehydrogenases.
DOI: https://doi.org/10.7554/eLife.33953.029
Figure supplement 4. Cgr2 is a distinct flavoprotein reductase.
DOI: https://doi.org/10.7554/eLife.33953.030

these results suggest that Cgr2 activity is restricted to cardenolide toxins and does not extend toother structurally related endogenous or exogenous compounds.
Additionally, neither fumarate nor any of the metabolized cardenolides conferred a growthadvantage to cgr2+ E. lenta in minimal or rich medias, suggesting that these compounds are notused as alternative terminal electron acceptors (Figure 4—figure supplement 2). The inability of
Cgr2 to reduce fumarate, a common electron acceptor used during bacterial anaerobic respiration,
led us to revisit the original annotation of Cgr2 as a ‘fumarate reductase’ (Saunders et al., 2009). To
more systematically assess the relationship between Cgr2 and biochemically characterized reductases, we constructed a sequence similarity network (SSN) using the 5000 most similar sequencesfrom the UniProtKB protein database. Within the network, there were seven enzymes that had been
biochemically characterized (UniProtKB IDs: Q07WU7, Q9Z4P0, 8CVD0, P71864), biochemically andstructurally characterized (PDB IDs: 1D4D, 1E39), or genetically characterized (UniProtKB ID:
Q7D5C1) (Figure 5—source data 1) (Leys et al., 1999; Bogachev et al., 2012; Brzostek et al.,2005; Doherty et al., 2000; Knol et al., 2008; Rothery et al., 2003; Pealing et al., 1992;
Dobbin et al., 1999). At all thresholds at which Cgr2 remained connected to other protein sequences, all characterized enzymes within the SSN were co-clustered, precluding the resolution of uniquebiochemical functions at this cutoff (Figure 5A). At higher alignment thresholds that separated thesecharacterized enzymes into discrete isofunctional clusters, Cgr2 was always present as a ‘singleton’with no links to other protein sequences (Figure 5B). Our SSN also revealed that reductase enzymes
are widespread among human gut bacteria, with three validated enzymatic activities and 113 distinct
clusters with uncharacterized biochemical functions detected in sequenced gut bacterial genomes.
We validated our SSN by aligning the biochemically characterized enzymes within the networkwith additional co-clustered sequences to assess conservation of essential active site residues (Figure 5—figure supplements 1–3). Comparing the sequence of Cgr2 with sequences of biochemicallycharacterized reductases revealed that Cgr2 lacks active site residues required for the activity offumarate reductases (6/7 divergent residues), urocanate reductases (4/5 divergent residues), andketosteroid dehydrogenases (3/5 divergent residues) (Figure 5—figure supplement 4A–D)(Leys et al., 1999; Rohman et al., 2013; Bogachev et al., 2012; Knol et al., 2008; Reid et al.,2000). Individually mutating the two residues shared between Cgr2 and ketosteroid dehydrogenases confirmed that one amino acid involved in substrate binding (G536 backbone) was also important for the activity of Cgr2 but the other (Y532) was not (Figure 5—figure supplement 4E).
Together, the location of Cgr2 within the SSN and the differences in its sequence indicate that thisenzyme is distinct from characterized bacterial reductases and may use a unique set of residues tocatalyze cardenolide reduction.

Cgr2 is widespread in the human gut microbiome
To assess the broader relevance of this cardenolide-metabolizing enzyme, we quantified the prevalence, conservation, and genomic context of cgr2 in the human gut microbiome. We mined gut

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Figure 6. Cgr2 is widespread in the human gut microbiome. (A) Analysis of the cgr-associated gene cluster and E. lenta (via elenmrk1) prevalence inthe gut metagenomes of 1872 individuals (see Materials and methods) revealed that both E. lenta and cgr2 are highly prevalent (41.5% and 27.7%
respectively) but frequently low in abundance. (B) Quantification of E. lenta and cgr2 abundances in individual gut metagenomes revealed a tightcorrelation between the two, providing evidence that cgr2 is restricted to E. lenta and that individuals may harbor sub-populations of both cgr2+ andcgr2- strains. Red line denotes the expected linear relationship and dashed lines represent a ± half log deviation. (Inset) Histogram of cgr-ratio (cgr/elnmrk1) demonstrates a significant skew away from communities that would have more cgr2 than expected by E. lenta abundance (p<0.001,
D’Agostino skewness test). (C) Replication in an additional 158 individuals located in the USA (n = 85) and Germany (n = 73) via duplexed qPCR
increased prevalence estimates to 74.7% and 81.6% at the extremes of detection limit (1e3 copies/g). qPCR samples were run in technical triplicate(Figure 6—source datas 1 and 2). (D) Similarly, qPCR-derived abundances of E. lenta and cgr2 were correlated, corroborating metagenome-based
analysis. (Inset) Histogram of cgr-ratio demonstrating significant skew (p<0.001).
DOI: https://doi.org/10.7554/eLife.33953.032
The following source data is available for figure 6:
Source data 1. qPCR based E. lenta and cgr2 prevalence.
DOI: https://doi.org/10.7554/eLife.33953.033
Source data 2. Replicate qPCR data of for cgr2 and elnmrk1 in human fecal samples.
DOI: https://doi.org/10.7554/eLife.33953.034

microbiome datasets from 1872 individuals sampled in 6 countries across three continents(Nayfach et al., 2015). Analysis of the E. lenta pan-genome led to the discovery of a single copy
marker gene (referred to here as elnmrk1) conserved in all sequenced strains that serves as a proxyfor E. lenta abundance in both sequencing and quantitative PCR (qPCR) assays (Bisanz et al., 2018).
The abundance of elnmrk1 was significantly associated with E. lenta abundance across the

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Microbiology and Infectious Diseaseindividuals (R2 = 0.973, p<2.2e-16). Using this marker gene, we detected E. lenta in 41.5% of subjects at abundances of 3.5 ± 0.58 mean ± SD log10(elnmrk1 copies/cell) (Figure 6A).
Cgr2 was detectable in 48.5% of the E. lenta-positive individuals, while detection occurred in
27.7% of all subjects ( 3.7 ± 0.64 mean ± SD log10(cgr2 copies/cell)) (Figure 6A), and the abundanceof E. lenta and cgr2 was significantly associated (rho = 0.455, p<0.001; Figure 6B). The distributions
were skewed towards participants with less cgr2 than expected based on the abundance of E. lenta[(skew = 0.773, p=2.5e-8, n = 375, D’Agostino skewness test of log10(cgr/elnmrk1) with quantifiable elnmrk1 and cgr2], consistent with prior data suggesting that many individuals are colonized bya mixture of cgr2+ and cgr2- E. lenta strains (Haiser et al., 2013). These results were validated by
qPCR in an independent set of 158 individuals (228 samples) from multiple sites in the USA and Germany (Figure 6—source data 1) revealing a similar skew towards higher E. lenta abundances versus
cgr2 (skew = 0.65, p<0.001, n = 165). Using this more sensitive detection method, we detected E.
lenta in 81.6% of individuals (1.5e7 ± 3.5e6 copies/g feces) and cgr2 in 74.7% (1.5e6 ± 3.5e6 copies/
g feces) (Figure 6C–D). Similar to the sequence-based analysis, the outliers were skewed towards

Figure 7. Preliminary model for digoxin metabolism by Cgr1 and Cgr2. (A) Proposed biochemical model and (B) mechanism of digoxin reduction by
Cgr proteins. Cgr1 is predicted to transfer electrons from a membrane-associated electron donor to the [4Fe-4S]2+ cluster of Cgr2 via covalently boundheme groups. The reduced [4Fe-4S]1+ cluster of Cgr2 could sequentially transfer two electrons to FAD, generating FADH–, which could mediate
hydride transfer to the b-position of the digoxin lactone ring. Protonation of the resulting intermediate would yield (20R)-dihydrodigoxin.
DOI: https://doi.org/10.7554/eLife.33953.035

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samples with less cgr2 than expected based on the abundance of E. lenta. Overall, both the qPCRand metagenomic sequencing-based analyses show that E. lenta and cgr2 are widely distributed in
the human microbiome.

Discussion
For over three decades, the human gut bacterium E. lenta has been linked to cardiac drug inactivation (Saha et al., 1983). However, the identity, specificity, and distribution of the enzymes responsible for this activity were unknown. In this work, we unambiguously show that the E. lenta protein,Cgr2, inactivates cardenolides, including the pharmaceutical agents digoxin and digitoxin that havebeen used for over two centuries in the treatment of cardiac diseases. Although we cannot rule out
the possibility that other enzymes are involved in digoxin inactivation, no other microbes have been
discovered that possess this metabolic activity apart from cgr2+ E. lenta (Saha et al., 1983). Cgr2
represents a novel flavoprotein reductase, and contains oxygen-sensitive [4Fe-4S] cluster(s) and adivergent set of predicted active site residues. The failure of bioinformatic analyses to identify thisessential [4Fe-4S] cluster highlights the need for additional structural and mechanistic studies of thecgr operon and other gut microbial enzymes involved in xenobiotic metabolism. Our working modelis that Cgr1 and Cgr2 form a membrane-anchored, extracellular complex that mediates electrontransfer from an electron donor (e.g. the membrane quinone pool) through multiple cytochromes cin Cgr1 to Cgr2, which ultimately reduces the a,b-unsaturated g-butyrolactone of digoxin and othercardenolides (Figure 7A). We propose that the [4Fe-4S]2+ cluster(s) of Cgr2 sequentially transfer
electrons to the FAD cofactor. The resulting hydride equivalent is then transferred to the cardenolide, and proton transfer generates the fully reduced g-butyrolactone (Figure 7B), yielding the therapeutically inactive metabolite dihydrodigoxin.
While we have demonstrated that Cgr2 is necessary and sufficient for digoxin reduction in a heterologous host and in vitro using a chemical electron donor, additional proteins within the expanded
cgr gene cluster may be important for digoxin reduction in vivo. Sequence analyses and transcriptional data suggest that Cgr1 is likely important for this metabolic activity in E. lenta. However, wewere unable to observe overexpression or heme c incorporation into Cgr1 using a variety of heterologous constructs, hosts, and expression conditions, which may be due to an incompatibility of heterologous cytochrome c maturation factors and this protein (Sanders and Lill, 2000; ThönyMeyer, 2002). The use of alternative heterologous systems that are more suitable for producing
multi-heme cytochromes c (Ozawa et al., 2001; Kern and Simon, 2011) or the development ofgenetic tools in E. lenta would thus be required to obtain functional Cgr1 and determine its role indigoxin metabolism.
In contrast to Cgr1, the relevance of the Cac proteins to the Cgr proteins and digoxin metabolism
is unclear. Apart from the putative LuxR-type regulator Cac3, which was modestly upregulated in
response to digoxin and may be involved in regulating transcription of the cgr operon, RNAsequencing does not show clear evidence for the expression of the other cac genes during growthin pure culture. While Cac4 is annotated as a secreted, FAD-dependent fumarate reductase, it lacks
all known catalytic and binding residues for this enzyme class (Figure 5—figure supplement 1). Furthermore, Cac4 shares only 23% sequence identity to Cgr2, and is thus likely to metabolize a different substrate than either of these enzymes. Cac6 is homologous to stomatin, prohibitin, flotillin, and
HflK/C (SPFH) proteins, which are often associated with lipid rafts or functional microdomains in bacteria (Bramkamp and Lopez, 2015) and could potentially interact with the Cgr enzymes, the substrates of the cgr operon, or additional steroidal substrates of E. lenta (Ridlon et al., 2006;
Devlin and Fischbach, 2015). Additional work is required to understand the biochemical function of
the Cac proteins and whether they influence E. lenta metabolism of digoxin and/or other small
molecules.
We have demonstrated that E. lenta strains harboring the cgr operon are widespread in thehuman gut microbiome, which supports the high incidence of dihydrodigoxin production observed
clinically. Strikingly, cgr2 and its associated genes are highly conserved, with two naturally occurring
variants (frequent: Y333/N333; infrequent: V381/M381). This conservation is surprising given thestrict specificity of Cgr2 towards cardenolides, which are ingested at very low concentrations to minimize toxicity in the context of cardiac therapy (Smith et al., 1969; Gheorghiade et al., 2004). These
results, together with the overall high degree of conservation in the E. lenta pan-genome, suggest

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that much of the phenotypic variation within this species may be driven by gene gain/loss rather
than by genetic polymorphisms. Data from other bacterial lineages suggests that this phenomenon
may not be unique to Coriobacteriia (Hao and Golding, 2006; Marri et al., 2006; Marri et al.,2007; Nowell et al., 2014; Vos et al., 2015); in Pseudomonas syringae, 1% amino acid divergenceaccumulates at the same time in hundreds or even thousands of genes (Nowell et al., 2014).
The high sequence conservation and levels of cgr operon transcription in response to digoxin
exposure suggest that digoxin metabolism may provide a physiological benefit to E. lenta. However,as we could not observe any direct benefit of cardenolide metabolism for cgr+ E. lenta (Figure 4—
figure supplement 2), we hypothesize that these bacteria may have evolved to protect the hostfrom plant toxins and thus maintain a habitat for colonization. Although we cannot rule out an as-ofyet unidentified endogenous substrate of Cgr2, our results suggest that similar to intestinal andhepatic enzymes, gut bacteria have co-evolved with the human host and maintain detoxification systems that can be rapidly and efficiently mobilized on demand. Additional studies are warranted that
directly compare and contrast the gut microbiomes of herbivorous animals and insects that evolved
under more constant exposure to cardenolides (Agrawal et al., 2012), as these communities may
represent a reservoir for evolutionary or functional homologs of cgr2. While we do not yet understand the factors that maintain this gene in the E. lenta population in the absence of a direct selective pressure, our studies, coupled with ex vivo experiments with human fecal samples and in vivo
experiments in mice (Haiser et al., 2013), suggest that the metabolic activities of low abundance
members of the gut microbiome can significantly influence host physiology.
Our results also highlight an important consideration for ongoing efforts to predict and manipulate gut microbial metabolism, particularly in the context of therapeutics. Not only do E. lenta strains
vary in the presence or absence of cgr2, but we have also identified a naturally occurring, singleamino acid substitution that causes a dramatic loss of activity in the Cgr2 enzyme. This result emphasizes the need for methods that can achieve nucleotide-level precision in mapping inter-individual
differences in human gut microbiome gene content. Several clinical trials have recently investigated
the use of digoxin for treating diverse cancers (Lin et al., 2014; Kayali et al., 2011), rheumatoid
arthritis (Huh et al., 2011) and HIV-1 infection (Wong et al., 2013). Paired studies of host genetics
combined with nucleotide-resolution analyses of the gut microbiome are needed to test the feasibility of using microbial genetic information to predict drug bioavailability and improve treatment outcomes in these new disease contexts.
Finally, our results demonstrate the feasibility of progressing from case studies of a clinically relevant gut microbial biotransformation (Saha et al., 1983) to identifying the responsible genes,
enzymes, and biochemical mechanisms associated with metabolism. Isolation of individual xenobiotic
metabolizing strains is a crucial first step towards uncovering the genetic and biochemical bases ofthese transformations. Starting from complex microbial communities (e.g. human fecal samples),individual strains can be selectively enriched (Kumano et al., 2016; Martı́nez-del Campo et al.,2015) or isolated (Saha et al., 1983; Bisanz et al., 2018) to enable screening and identification of
microbes that metabolize a xenobiotic of interest. The observation that many xenobiotic-processing
or transporting genes are only upregulated in the presence of substrate can be leveraged to identify
xenobiotic-metabolizing genes using techniques such as RNA-seq (Haiser et al., 2013) or native protein purification (Kumano et al., 2016). Genetic and/or heterologous expression experiments canthen help to validate the role of the identified genes and enzymes in xenobiotic transformation.
Finally, the responsible genes can serve as candidate biomarkers to probe the distribution andpotential for xenobiotic metabolism in ex vivo incubations or in relevant clinical populations.
These studies also provide new insights into the chemistry made possible by complex host-associated microbial communities. The gut microbiome encodes over 3 million genes (Qin et al., 2010)
and >50% have unknown functions (Human Microbiome Project Consortium, 2012). Our study ofjust one highly unique, clinically relevant gut microbial enzyme has illuminated underappreciatedfunctional diversity within the broader flavin-dependent reductases, which is widespread among
human gut microbes (Figure 5). As reductive transformations represent a major route by which gut
microbes metabolize xenobiotics, these unique, putative reductase enzymes provide a promising
starting point for identifying additional gut microbiome-xenobiotic interactions. The approaches
described here, together with high-throughput methods to characterize protein subfamilies andadvances in bacterial culturing and genetic tools (Goodman et al., 2011; Lim et al., 2017), are

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beginning to unlock the genetic ‘dark matter’ of the human gut microbiome as well as its critical rolein the etiology and treatment of human disease.

Materials and methods
Key resources table
Reagent type (species)
or resource

Designation

Source or reference

Identifiers

Additional information

Gene(Eggerthella lenta)

cgr1

PMCID: PMC3035228

cgr1

Gene
(Eggerthella lenta)

cgr2

PMCID: PMC3035228

cgr2

Gene
(Eggerthella lenta)

16S rRNA

Ref 11

Gene(Eggerthella lenta)

E. lenta marker geneRef 11

Strain, strain
background(Escherichia coli)

One Shot Top10

Thermo Fisher Scientific

Strain, strain
background(Rhodococcus
erythropolis)

L88

doi: 10.1128/JB.
187.8.2582–2591.2005

Strain, strain
background(Eggerthella lenta)

Eggerthella lenta
strains

Ref 11El1 - El21

Strain, strain
background(Eggerthella sinensis)

Eggerthella sinensis DSM16107

Ref 11

Es1

Strain, strain
background(Gordonibacter)

Gordonibacter strains

Ref 11

Gs1, Gs2

Strain, strain
background(Paraeggerthella
hongkongensis)

Paraeggerthella hongkongensis

Ref 11

Ph1

Recombinant
DNA reagent

pTip expression
vectorsdoi: 10.1128/AEM.70.9.5557–5568.2004

Sequence-based
reagent

Amplification cgr1,
cgr2

Integrated DNA
technologies

see Table 1 for primers
and constructs

Sequence-based
reagent

Point mutants of
Cgr2

Integrated DNA
technologies

see Table 2 for primers
and mutants

Sequence-based
reagent

cgr2 sequencing
primerThis work

Confirmed sequence of E. lenta
isolates with primers: cgr2_fwd(TGCAATCAAGACAACCACGA),
cgr2_internal (TCGGTGTACAACCACAATGC),
and cgr2_rev (GTTGCGCTGTGATTAGACTG)

Sequence-based
reagent

qPCR primers cgr2This work

cgr2_F (GAGGCCGTCGATTGGATGAT),
cgr2_R (ACCGTAGGCATTGTGGTTGT),and cgr2_probe ([HEX]CGACACGG
AGGCCGATGTCG[BHQ1])

Sequence-based
reagent

qPCR primers elnmrk

This work

ElentaUni_F (GTACAACATGCTCCTTGCGG),
ElentaUni_R (CGAACAGAGGATCGGGATGG),
ElentaUni_Probe ([6FAM]TTCTGGCTGC
ACCGTTCGCGGTCCA[BHQ1]),

Chemicalcompound, drug

BBL Brain Heart
Infusion (BHI) media

Becton Dickinson

BD:L007440

Chemicalcompound, drug

L-arginine

Sigma Aldrich

SA:A5006

elnmrk1

see Figure 1—source data 1
for full descriptions

Continued on next page

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Continued
Reagent type (species)
or resource

Designation

Source or reference

Identifiers

Chemicalcompound, drug

Digoxin

Sigma Aldrich

SA:D6003

Chemicalcompound, drug

Dihydrodigoxin

doi: 10.1126/
science.1235872

Chemicalcompound, drug

Digitoxin

Sigma Aldrich

SA:D5878

Chemicalcompound, drug

Digoxigenin

Sigma Aldrich

SA:D9026

Chemicalcompound, drug

Ouabain

Sigma Aldrich

SA:O3125

Chemicalcompound, drug

Ouabagenin

Sigma Aldrich

SA:O2627

Chemicalcompound, drug

Sypro Orange
protein gel stain

Thermo Fisher Scientific

SA:S6650

Chemicalcompound, drug

Thiostrepton

Sigma Aldrich

SA:T8902

Chemicalcompound, drug

Methyl viologen

Sigma Aldrich

SA:856177

Chemicalcompound, drug

Sodium dithionite

Sigma Aldrich

SA:157953

Chemicalcompound, drug

FAD

Sigma Aldrich

SA:F6625

Chemicalcompound, drug

Iron (II) ammonium
sulfate hexahydrate

Sigma Aldrich

SA:F1543

Chemicalcompound, drug

Sodium sulfide nonahydrate

Sigma Aldrich

SA:208043

Chemicalcompound, drug

Dithothreitol

Sigma Aldrich

SA:D0632

Chemicalcompound, drug

HEPES

Sigma Aldrich

SA:RDD002

Software,algorithm

Cytoscapedoi:10.1101/gr.1239303

Software,algorithm

Prism software

Prism software

Other

Anaerobic chambers

Coy Laboratory
products; Mbraun

Other

LC-MS/MS

Agilent

Other

Electron paramagentic
resonance (EPR)
spectrometer

BrukerOther

CFX96 Touch RealTime PCR machine

Bio-Rad

Other

PowerWave HT
Microplate Spectrophotometer

BioTekAdditional information

Graphpad
Software v 7

Agilent:6410
TripleQuad LC/MS

Genome analysis
Publically available genomes were retrieved from NCBI (E. lenta DSM 2243, PRJNA21093; E. lenta
FAA1-3-56 PRJNA40023). New isolates were sequenced as described elsewhere (Bisanz, et al.
2018). Genomes were assembled with SPAdes 3.11.1 (84) and annotated with Prokka 1.12 (Seemann, 2014). All E. lenta strains studied were identified as E. lenta based on 16S rRNA sequencing
and were de-replicated at the strain level by considering a pairwise average nucleotide identity(ANI) >99.99% as the same strain (github.com/widdowquinn/pyani). The maximum and minimum

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ANI between studied E. lenta strains were 97.9% and 99.6% respectively. The phylogenetic tree wasprepared using a set of 400 conserved proteins (Segata et al., 2013) rooted on the Gordonibacterstrains. Newly sequenced strains were included as part of Bioproject PRJNA412637.
Global nucleotide and amino acid identity and related statistics were determined via the Needleman-Wunsh implementation in the pairwiseAlignment function of Biostrings with percentage identity
calculated as 100*identical positions/(aligned positions + internal gaps). Statistical and graphicalanalysis was carried out using R 3.4.0. RNA sequencing reanalysis was carried out by mapping reads
to the reference genome with Bowtie2 (Langmead and Salzberg, 2012), counting with HTSeq(Anders et al., 2015), and differential expression analysis using DESeq2 (Love et al., 2014). Original
sequence data is available from the SRA with project identifier SRP018311.
For the purposes of comparative genomics, gene conservation was calculated by first clustering
into orthologous clusters with proteinortho5 (Lechner et al., 2011) with a minimum 60% amino acid
identity and 80% coverage. A presence/absence matrix de-replicated for co-occurring features wasthen used as the input for a random forest classifier (randomForest 4.6–12). Variable importance(mean decrease GINI) was used to extract the 15 most important features. A tool for this comparative genomic analysis is available as ElenMatchR (jbisanz.shinyapps.io/elenmatchr; copy archived athttps://github.com/elifesciences-publications/ElenMatchR) (Bisanz and Turnbaugh, 2018) withdigoxin reduction available as a demonstration dataset.

Bacterial culturing
Eggerthella lenta and related strains were grown in BBL Brain Heart Infusion (BHI) media (BD, Franklin Lakes, NJ) supplemented with L-arginine (Sigma-Aldrich, St. Louis, MO) under an atmosphere of
2–5% H2, 2–5% CO2, and balance N2. Strains were streaked onto BHI agar plates supplemented
with 1% arginine (w/v) in an anaerobic chamber (Coy Laboratory Products, Grass Lakes, MI). Individual colonies were inoculated into 16  125 mm Hungate tubes (Chemglass Life Sciences, Vineland,NJ) containing 5–10 mL of BHI supplemented with 1% arginine and grown at 37˚C for 2–3 days. Cardiac glycoside substrates were dissolved at a concentration of 10 mM in dimethylformamide (DMF)
and added to cultures at a final concentration of 10 mM. Starter cultures were diluted into 10 mL of
BHI + substrate to a starting of OD600 of 0.05 and grown anaerobically at 37˚C for 2 days. Experiments were performed in biological triplicate.
For growth assays, E. lenta DSM 2243 was grown in either rich (BHI) or defined media. Basal
media lacking terminal electron acceptors was prepared as previously described (Löffler et al.,2005) with the following modifications: yeast extract and tryptone were each added to 0.1% (w/v),
L-cysteine concentration was 0.4 mM, sodium sulfide was not added, and either 5% H2 or 10 mM
sodium acetate were used as electron donors. Starter cultures were prepared as described above in
BHI media supplemented with 1% arginine, and diluted 1:100 into media that had been supplemented with substrates (dissolved in DMF) to a final concentration of 10 mM. Cultures were grownanaerobically at 37˚C in biological triplicate. OD600 measurements were recorded on a Genesys20
spectrophotometer (Thermo Fisher Scientific, Waltham, MA).

Extraction and LC-MS/MS detection of digoxin and dihydrodigoxin
Bacterial cultures were centrifuged (10 min x 4000 rpm), 1 mL of supernatant was extracted threetimes with 1 mL of dichloromethane and the pooled organic fractions were concentrated using arotary evaporator. Samples were resuspended in 1 mL of 50% methanol in water and diluted 10xprior to liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis.
Metabolites were detected on an Agilent 6410 Triple Quad LC/MS using electrospray ionizationin negative ion mode. The mass spectrometer settings were as follows: gas temperature (300˚C), gas
flow (10 L/min), nebulizer pressure (25 psi), capillary voltage (4000 V), and chamber current (0.1 mA).
Digoxin was monitored using a 779.4 fi 649.3 m/z transition with a fragmentor voltage of 250V andcollision energy of 52, and dihydrodigoxin was monitored using a 781.4 fi 521.3 m/z transition witha fragmentor voltage of 200V and collision energy of 20. Standard curves were prepared using 0.01–
1 mM samples of each compound. Digoxin was purchased from Sigma-Aldrich (St. Louis, MO), and adihydrodigoxin standard was obtained through chemical hydrogenation of digoxin as previously
described (Haiser et al., 2013). Liquid chromatography was performed on an Acclaim Polar Advantage II column with a flow rate of 0.125 mL/min, 5 mL sample injection, solvent A (10% methanol + 1

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Table 1. Primers and constructs for heterologous expression of Cgr proteins in R. erythropolis.
Restriction sites are bolded.
For/
Constructcgr operon

Rev Sequence

Restriction
sites
VectorAnneal Extend
temp
time
(˚C)
(s)

For

ACTGACCCATGGATGGAATACGGAAAGTGCC

n/a

n/a

71

75

Rev

GTTTTACTGCAGTTACGCCGCCGTCGAATGACGAATTCTAATGGAATACGGAAAGTGCCG

EcoRI,

pTipQT2 70

90

Rev

TTATAAGATCTCGCCGCCGTCGAAAG

BglII

For

TCGAACATATGATGGCTGAGGAACCTGTGG

NdeI,

pTipQT1 65

60

Rev

ATAACTCGAGTCACGCCGCCGTCGAAA

XhoI

ACTGACCCATGGGCATGGAATACGGAAAGTGCC NcoI,
HindIIIATTAGAAGCTTTCACTCCCACGGCTCGAG

pTipQC2 65

60

pTipQC1 65

60

NcoI,
HindIII

pTipQC2 65

60

NcoI,
HindIII

pTipQC1 65

60

n/a

All pTip
vectorsn/a

Cgr1 + Cgr2 For

Cgr1

Cgr2 (native) For
Rev
Cgr2-CHis6(native)

For
Rev

ACTGACCCATGGGCATGGAATACGGAAAGTGCC NcoI,
HindIIIGTTAGAAGCTTCTCCCACGGCTCGAG

Cgr2( 48aa)NHis6

For

TATTACCATGGATCAGACCGCGCCTGC

Rev

ATACTAAGCTTCTCCCACGGCTCGA

Cgr2( 48aa)CHis6

For

TATTACCATGGATCAGACCGCGCCTGC

Rev

ATACTAAGCTTTTACTCCCACGGCTCGA

Sequencing
primers

For

CGTGGCACGCGGAAC

Rev

GTGCAGGTTTCGCGTG

n/a

DOI: https://doi.org/10.7554/eLife.33953.036

mM ammonium hydroxide) and solvent B (100% methanol + 1 mM ammonium hydroxide), and a
gradient: 70–100% B over 10 min, 100% B for 1.5 min, 100–70% B over 3.5 min, and 70% B for 7
min.

Construction of cgr1 and cgr2 vectors in Escherichia coli
E. lenta DSM 2243 was grown in 5 mL of BHI + 1% arginine at 37˚C. After 2 days, the culture waspelleted and genomic DNA (gDNA) was purified with the UltraClean Microbial DNA Isolation Kit(QIAGEN, Germantown, MD) according to the manufacturer’s protocol. The cgr operon was amplified from 50 ng of gDNA in a 50 mL reaction volume with 0.5 mM of each primer (Table 1) and Phusion High-Fidelity PCR master mix with HF buffer (New England Biolabs, Ipswich, MA). The following
thermocycling parameters were used: denaturation at 98˚C for 3 min; 35 cycles of 98˚C for 15 s,
71˚C for 20 s, and 72˚C for 75 s; and a final extension at 72˚C for 5 min. The PCR reaction was analyzed by agarose gel electrophoresis (1% agarose gel), and the cgr amplicon was excised and purified with the Illustra GFX PCR DNA and Gel Band Purification kit (GE Healthcare, Chicago, IL). Cgr1
and cgr2 variants were amplified in 20 mL PCR reactions using 1 ng of purified cgr operon as template, 0.5 mM primer pairs and Phusion High-Fidelity PCR master mix with HF buffer (New England
Biolabs, Ipswich, MA) (Table 1). PCR conditions were as follows: denaturation at 98˚C for 2 min; 35
cycles of 10 s at 98˚C, 10 s at specified annealing temperature, and 72˚C for the specified extension
time; and a final extension at 72˚C for 5 min. Cgr amplicons were digested in a 30 mL reaction with
1.5 mL of each restriction enzyme (New England Biolabs, Ipswich, MA) for 2.5 hr at 37˚C. pTip vectors were similarly digested, and the linearized vector was excised from a 1% agarose gel and purified. Insert and vector pairs were ligated at a 1:3 ratio at room temperature for 2 hr with T4 DNA
ligase (New England Biolabs, Ipswich, MA). 2.5 mL of the ligation reaction was transformed into
chemically competent One Shot Top10 E. coli cells (Thermo Fisher Scientific, Waltham, MA) andplated on LB agar with ampicillin. Plasmid inserts were sequenced using the primers listed inTable 1.

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Table 2. Primers for site-directed mutagenesis of Cgr2.
Amino acid numbering is based on full length Cgr2 sequence. Introduced mutations are bolded.
Mutant
C82A

C111A

C158A

C187A

C231A

F/R

SequenceFor

CAGCGGCGGCACGGCCGCGGCCATCG

Rev

CCTCGATGGCCGCGGCCGTGCCGCCGFor

GCGGCAACTCGGCACTAGCCGGTGGATACATRev

CCAGCATGTATCCACCGGCTAGTGCCGAGTTG

For

ATATGATCCGCGAGGCGGCCTTGCGCTCCGGC

Rev

GCCTCGCCGGAGCGCAAGGCCGCCTCGCGGATFor

GCCCCCGGTCTGGTCAGCCGGCGACACGG

Rev

GGCCTCCGTGTCGCCGGCTGACCAGACCGGForCGAAATCGAGATGGGCGCCGAGGTGGCGCAC

Rev

GATGTGCGCCACCTCGGCGCCCATCTCGAT

C265A

For

GGCGTGGTCATGGCGGCCGCTTCGGTGGARev

GTTGTCCACCGAAGCGGCCGCCATGACCA

C321A

For

GATCGGTGCTGAGCTTGCCATGCAGCAGGC

Rev

CACGGCCTGCTGCATGGCAAGCTCAGCACC

For

CATGCAGCAGGCCGTGGCCATGAACGATTCTRevGATAGAATCGTTCATGGCCACGGCCTGCTG

For

GACCGGCAGACGGTTTGCCCAGGACGATGCCG

Rev

CTCGGCATCGTCCTGGGCAAACCGTCTGCC

For

CTATGTCATGCACGAGGCCGCGCAAGCTGCA

Rev

CCATGCAGCTTGCGCGGCCTCGTGCATGAC

C327A

C371A

C384A

C425A

C443A

C459A

C483A

C521A

For

CATACGCCCGACACGGCCGATACTACGTTCRevCGAGAACGTAGTATCGGCCGTGTCGGGCGTFor

GCCGAGTTTATCGGCGCCGATCCGACCGC

Rev

GAGGGCGGTCGGATCGGCGCCGATAAACTCFor

GAGGTGGAACTCTTTCGCCGAGGCCGGTTTG

Rev

CATCCAAACCGGCCTCGGCGAAAGAGTTCCAForGACGCCGCCGTTCTACGCCGATGTCGTGCGC

Rev

GGGGCGCACGACATCGGCGTAGAACGGCGG

For

CTGTACGGCGCCGGGGCCATCATCGGGGGTRev

GTTACCCCCGATGATGGCCCCGGCGCCGTA

C535A

For

GCCTTCTACTTCGGCGCCGGCTGGTCCATC

Rev

CGTGATGGACCAGCCGGCGCCGAAGTAGAA

Y333N

For

GCATGAACGATTCTATCAACGTAGGCGGCATCA

Rev

TCGCTGATGCCGCCTACGTTGATAGAATCGTTCA

Y532F

For

GATGCCGAGTGGGGCTTTGTCATGCACG

Rev

GCACTCGTGCATGACAAAGCCCCACTCG

G536A

For

TTCTACTTCGGCTGCGCCTGGTCCATCARev

GTTCGTGATGGACCAGGCGCAGCCGAAG

DOI: https://doi.org/10.7554/eLife.33953.037

Site-directed mutagenesis of Cgr2
Site-directed mutagenesis was performed in 25 mL reactions using 200 ng of template DNA (Cgr2(–48aa)-NHis6 in pTipQC2), 0.5 mM of each primer pair (Table 2), 0.5 mM dNTP, and 1 mL of Pfu Turbo

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polymerase AD (VWR, Radnor, PA). The following thermocycling parameters were used: denaturation at 95˚C for 1 min; 18 cycles of 95˚C for 30 s, 65˚C for 50 s, and 68˚C for 22 min (2 min/kb); anda final extension at 68˚C for 7 min. The template plasmid was digested with 1 mL of DpnI (New England Biolabs, Ipswich, MA) for 1 hr at 37˚C, and 2 mL of the reaction were transformed into chemically competent One Shot Top10 E. coli cells (Thermo Fisher Scientific, Waltham, MA).

Heterologous expression of Cgr proteins in Rhodococcus erythropolis
L-88
All Rhodococcus strains and expression vectors were obtained from the National Institute ofAdvanced Industrial Science and Technology (Tokyo, Japan). 40 ng of plasmid DNA were added to
400 mL of R. erythropolis L-88 electrocompetent cells in 30% PEG 1000 (Sigma-Aldrich, St. Louis,MO) in a 2 mm gap electroporation cuvette (VWR, Radnor, PA). Cells were transformed in a MicroPulser electroporator (Bio-Rad, Hercules, CA) with a 2.5 kV pulse (time constant ~4.8 – 5.2), rescued
with 0.6 mL of LB (Lennox) broth (Alfa Aesar, Tewksbury, MA), and incubated for 4 hr at 28˚C, 175
rpm. Cells were plated onto LB agar plates +antibiotic (17 mg/mL chloramphenicol for pTipQC plasmids; 8 mg/mL tetracycline for pTipQT plasmids) and incubated at 28˚C for 5–7 days. Single colonies
were inoculated into 50–75 mL of LB +antibiotic (34 mg/mL chloramphenicol or 8 mg/mL tetracycline)
and grown for 3–5 days at 28˚C, 175 rpm until reaching saturation. For gain of function studies, 50
mL of LB and antibiotic were inoculated to a starting OD600 of 0.2 and grown at 28˚C, 175 rpm.
Experiments were performed in biological triplicate. When cultures reached an OD600 of 0.6 (~6–8hr), protein expression was induced with thiostrepton (Sigma-Aldrich, St. Louis, MO) at a final concentration of 0.01 mg/mL, and cultures were incubated at 15˚C, 175 rpm. In cultures where Cgr1 wasoverexpressed, media was supplemented with the heme precursor d-amino levulinic acid hydrochloride (50 mg/mL final concentration) (Frontier Scientific, Logan, Utah). After 16–20 hr, digoxin wasadded to cultures as a solution in DMF at a final concentration of 10 mM and incubated for either 7days at 15˚C, or 2 days at 28˚C, 175 rpm. Culture supernatants were extracted and analyzed as previously described. For large-scale purifications, 2 L of LB-chloramphenicol in a 4 L baffled flask were
inoculated to a starting OD600 of 0.02 and grown to an OD600 of 0.6 (~18–25 hr). Protein expression
was induced with 0.01 mg/mL thiostrepton, and cultures were incubated at 15˚C, 175 rpm forapproximately 21 hr before harvesting cells by centrifugation (10,800 rpm x 20 min). Cell pellets
were frozen and stored at –80˚C.

Cgr2 purification and [Fe-S] cluster reconstitution
All protein purification steps were carried out at 4˚C. Harvested cells were resuspended in 5 mL/g ofcell pellet in lysis buffer (50 mM Tris, pH 8, 1 mM MgCl2, 25 mM imidazole) containing Pierce EDTAfree protease inhibitor cocktail (Thermo Fisher Scientific, Waltham, MA). Cells were passaged
through a cell disruptor (Avestin EmulsiFlex-C3) five times at 15,000–25,000 psi and centrifuged for
20 min at 13,000 rpm. The clarified lysate was incubated on a nutating mixer with 5–10 mL of HisPur
Ni-NTA resin (Thermo Fisher Scientific, Waltham, MA) for 1 hr and then applied to a gravity flow column. The resin was washed with 50 mL of wash buffer (25 mM HEPES, 0.5 M NaCl, pH 8, 25 mM
imidazole) and eluted with 25 mL of elution buffer (25 mM HEPES, 0.5 M NaCl, pH 8, 200 mM imidazole). Eluted protein was concentrated using a 20 mL Spin-X UF 30 k MWCO PES spin filter (Corning, Corning, NY) to a volume of 1–2.5 mL, and then desalted on a Sephadex G-25 PD-10 desalting
column (GE Healthcare, Chicago, IL) that had been equilibrated with desalting buffer (50 mM
HEPES, 100 mM NaCl, pH 8). Desalted protein was sparged with argon on ice for 30–45 min. Chemical reconstitution of [Fe-S] cluster(s) in Cgr2 was carried out at 4˚C in an anaerobic chamber (CoyLaboratory Products, Grass Lakes, MI) under an atmosphere of 2% hydrogen and 98% nitrogen. A
30 mM solution of Cgr2 was prepared in reconstitution buffer (50 mM HEPES, 100 mM NaCl, pH 8,
and 2 mM dithiothreitol (DTT)). Fe(NH4)2(SO4)26H2O (Sigma-Aldrich, St. Louis, MO) was added infour aliquots over 60 min, followed by addition of Na2S9H20 (Sigma-Aldrich, St. Louis, MO) in four
aliquots over 60 min to final concentrations of 0.24 or 0.375 mM (8 or 12.5 equivalents relative toCgr2), and stirred for 16–24 hr. The reaction was filtered through a 0.25 mm, 0.2 mM pore-size PES
syringe filter (VWR, Radnor, PA) to remove precipitant and concentrated in a 6 mL Spin-X UF 30 k
MWCO PES spin filter inside a 50 mL conical-bottom centrifuge tube with plug seal cap (Corning,Corning, NY). The concentrated protein (1–2.5 mL) was desalted on a PD-10 column into desalting

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buffer. Protein was aliquoted into 0.5 mL PP conical tubes with skirt (Bio Plas), sealed inside 18 
150 mm Hungate tubes (Chemglass Life Sciences, Vineland, NJ) and stored at –80˚C. Protein concentration was determined by Bradford using Protein Assay Dye Reagent (Bio-Rad, Hercules, CA)and bovine serum albumin (BSA) (Sigma-Aldrich, St. Louis, MO) as a reference standard. Typical protein yields were ~20 mg/L of culture for both wild-type and point mutants of Cgr2(–48aa)-NHis6,~8
mg/L for Cgr2(–48aa)-CHis6, and ~1 mg/L for Cgr2-CHis6. The iron and sulfur content of Cgr2 samples (protein concentrations between 20–50 mM) was determined using previously reported colorimetric assays (Craciun et al., 2014).

Thermal denaturation assays
Thermal denaturation assays of purified and reconstituted Cgr2 were prepared on ice in 0.2 mL
skirted 96-well PCR plates (VWR, Radnor, PA) sealed with optical adhesive covers (Life Technologies,
Woburn, MA). Each reaction contained 10 mg of purified or reconstituted Cgr2, Sypro Orange protein gel stain (Thermo Fisher Scientific, Waltham, MA) diluted 5000-fold, and buffer containing 100
mM buffering agent and 100 mM NaCl in a total volume of 30 mL. The following buffering agentswere used: acetate/acetic acid for pH 4–6, HEPES for pH 7, Tris-HCl for pH 8–9, and glycine-NaOH
for pH 10. For metal binding assays, metal salts (Sigma-Aldrich, St. Louis, MO) were dissolved in pH
8 buffer to generate 100 mM stock solutions and added to a final concentration of 48 mM (8 equivalents relative to Cgr2). Data was collected on a CFX96 Touch Real-Time PCR machine (Bio-Rad, Hercules, CA) using the ‘FRET’ filter setting with FAM excitation and HEX emission channels (485 nm
and 556 nm respectively). The following temperature-scan protocol was used: 25˚C for 30 s, thenramp from 25˚C to 100˚C at a rate of 0.1 ˚C/ s.

Gel filtration
Gel filtration experiments were carried out on a Superdex 200 10/300 GL column (GE Healthcare,
Chicago, IL) attached to a BioLogic DuoFlow chromatography system (Bio-Rad, Hercules, CA).
Experiments were carried out either aerobically or anaerobically inside a Coy anaerobic chamber(Coy Laboratory Products, Grass Lakes, MI). 100 mL protein samples (50–100 mM) were loaded onto
the column at a rate of 0.2 mL/min for 1 mL followed by an isocratic flow of 0.33 mL/min for 30 mL
with 50 mM HEPES, 100 mM NaCl, pH 8. The molecular weight for Cgr2(–48aa)-NHis6 is 55.7 kDa. A
gel filtration standard (Bio-Rad, Hercules, CA) containing thyroglobulin (670 kDa), g-globulin (158kDa), ovalbumin (44 kDa), myoglobin (17 kDa), and vitamin B12 (1.35 kDa) was used to determine
the molecular weight of Cgr2-containing peaks.

UV-vis spectroscopy
Cgr2 was diluted to a final concentration of 50–100 mM in UV-Star UV-transparent 96-well microplates (Greiner Bio-One, Monroe, NC). The absorbance was measured between 250–750 nm using aPowerWave HT Microplate Spectrophotometer (BioTek, Winooski, VT) inside of an anaerobic glovebox (Mbraun, Stratham, NH). Curves were baseline subtracted using respective absorbance values at
700 nm. To determine whether the [Fe-S] cluster(s) were redox active, Cgr2 was incubated with 10
equivalents of sodium dithionite (Sigma-Aldrich, St. Louis, MO) for 15 min at room temperature prior
to taking additional absorption spectra. To assess the oxygen sensitivity of [Fe-S] cluster(s), Cgr2 wastaken out of the anaerobic chamber and exposed to oxygen, and the absorption spectra was measured aerobically on a PowerWave HT Microplate Spectrophotometer (BioTek, Winooski, VT). Oxygen-exposed Cgr2 was then sparged for 30 min with argon (on ice) and brought back into theMbraun glovebox for activity assays.

EPR spectroscopy
All samples were prepared in 50 mM HEPES, 100 mM NaCl, pH 8 under oxygen-free conditions in
an anaerobic glovebox (Mbraun, Stratham, NH). For all EPR experiments the final concentration ofCgr2 was either 150 or 200 mM. When required, the samples were reacted with an excess of sodium
dithionite (10–20 equivalents) for 20–30 min at 22˚C prior to freezing in liquid N2. Spin quantification
was carried out against a Cu2+-EDTA standard containing an equimolar concentration of CuSO4 in
10 mM EDTA (150 or 200 mM), under non-saturating conditions. Samples (250 mL) were loaded into
250 mm length, 4 mm medium wall diameter Suprasil EPR tubes (Wilmad LabGlass, Vineland, NJ)

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and frozen in liquid N2 under oxygen-free conditions. EPR spectra were acquired on a Bruker E500
Elexsys continuous wave (CW) X-Band spectrometer (operating at approx. 9.38 GHz) equipped witha rectangular resonator (TE102) and a continuous-flow cryostat (Oxford 910) with a temperature controller (Oxford ITC 503). The spectra were recorded at variable temperatures between 10–40 K at a
microwave power of 0.2 mW, using a modulation amplitude of 0.6 mT, a microwave frequency of
9.38 GHz, a conversion time of 82.07 ms, and a time constant of 81.92 ms.

In vitro substrate reduction assays
Methyl viologen (paraquat) dichloride hydrate (Sigma-Aldrich, St. Louis, MO) that had been reducedwith sodium dithionite was used as an artificial electron donor (Watanabe and Honda, 1982) to initiate anaerobic Cgr2-mediated reduction of digoxin in vitro. Assays were carried out at 25˚C in ananaerobic glovebox (Mbraun, Stratham, NH) under an atmosphere of nitrogen and < 5 ppm oxygen.
Reagents were brought into the glovebox as solids or sparged liquids and resuspended in anoxic
buffer inside the chamber: flavin (FAD or FMN) and methyl viologen (MV) were resuspended in 50
mM HEPES, 100 mM NaCl, pH 7 to generate stock solutions of 1 mM and 50 mM respectively;
sodium dithionite was resuspended in 50 mM HEPES, 100 mM NaCl, pH 8 to generate a stock solution of 25 mM; all substrates (Figure 4—figure supplement 1) were dissolved in DMF to generate
stock solutions of 25 mM, with the exception of sodium fumarate dibasic and urocanic acid whichwere dissolved in water. All substrates and reagents were purchased from Sigma-Aldrich (St. Louis,MO) except for the bufadienolides (Enzo Life Sciences, Farmingdale, NY) and prostaglandins (Cayman Chemicals, Ann Arbor, MI). The final assay mixture (100 mL) contained 5 mM Cgr2, 50 mM flavin,0.375 mM MV, 0.25 mM dithionite, and was initiated by addition of 0.5 mM substrate. For metal
activation studies, metal salts were dissolved in pH 7 buffer (1 mM) and added to a final concentration of 40 mM. Assays were prepared in a 96-well polysterene microplate (Corning, Corning, NY) andactivity was continuously monitored by measuring the absorbance at 600 nm on a PowerWave HT
Microplate Spectrophotometer (BioTek, Winooski, VT); a decrease in the absorbance at 600 nm corresponded to MV oxidation coupled to substrate reduction. For endpoint assay, reactions werequenched in methanol, diluted to a final concentration of 1 mM in 50% methanol, and analyzed by
LC-MS/MS as previously described.

Kinetic assays
Kinetic assays were performed in an anaerobic glovebox (Mbraun, Stratham, NH) at 25˚C. Reactions
were run in triplicate (200 mL) in assay buffer containing 5 mM Cgr2, 500 mM FAD, 1.5 mM MV, and 1
mM sodium dithionite, and were initiated by addition of digoxin as a solution in DMF to a final concentration of 0.01, 0.025, 0.05, 0.1, 0.2, 0.25, 0.3, and 0.5 mM. 20 mL reaction aliquots werequenched in 180 mL of ice-cold methanol in Costar flat bottom polysteryene 96-well plates (Corning,Corning, NY). The plates were sealed with adhesive aluminum foil for 96-well plates (VWR, Radnor,
PA), brought out of the anaerobic chamber, and further diluted (50-fold) into 50% methanol. Digoxin
and dihydrodigoxin standard curves were prepared in the full reaction matrix and processed identically such that final concentrations (after 500x total dilution) generated a standard curve between
0.01–1 mM. Plates were centrifuged (4000 rpm x 10 min, 4˚C) and 200 mL of each reaction weretransferred to a 0.5 mL PP 96-well plate (Agilent Technologies, Santa Clara, CA) sealed with EPS
easy piercing seals (BioChromato, San Diego, CA). The reactions were monitored by LC-MS/MS as
previously described, except that samples were directly injected (no column), and isocratic flow wasused with 75% methanol with 1 mM ammonium hydroxide.

Chemical similarity analysis
The chemical similarity of all substrates was assessed using the ChemMine software (http://chemminetools.ucr.edu) (Backman et al., 2011). Substrates were imported into ChemMine in SMILES format. The hierarchical clustering tool was used to generate a heatmap visualizing the structural
distance matrix between each substrate and digoxin.

Cgr2 sequence analysis
The full length Cgr2 protein sequence from E. lenta DSM 2243 was used as a query for BLASTP(Atlschul et al., 1997) using the NCBI non-redundant protein sequence database (search performed

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9/26/17). Cgr2 was also used to query the HHPred prediction tool (https://toolkit.tuebingen.mpg.
de/#/tools/hhpred) to identify additional remote protein homologs using hidden Markov models(Alva et al., 2016). The PDB_mmCIF70_27_Aug database was used (search performed 9/26/17).

Construction of sequence similarity network (SSN)
A SSN was generated using the EFI-EST tool (http://efi.igb.illinois.edu/efi-est/) (Gerlt et al., 2015).
The full length (native) Cgr2 protein sequence was used as an input to generate a network with the5000 most similar sequences from the UniProtKB protein database. An initial alignment score cutoffof 10 66 generated a SSN with 2018 nodes (with 100% identity) and 317,130 edges. The SSN was
imported into Cytoscope v 3.2.1 and visualized with the ‘Organic layout’ setting. Seven characterized enzymes were present within the network (UniProtKB IDs: fumarate reductases: P83223,
P0C278, Q07WU7, Q9Z4P0; urocanate reductase: Q8CVD0; 3-oxosteroid-1-dehydrogenase:P71864; Q7D5C1). The alignment score cutoff was increased to e-value <10 130, until enzymes withknown functions separated into putatively isofunctional clusters. At this threshold, Cgr2 appears as asingleton. The network shown in Figure 5A was generated with a cutoff of e-value <10 50, a threshold at which nearly all protein sequences form one cluster. Multiple sequence alignments were generated in Geneious and visualized in Jalview (clustalx coloring). To validate that the clusters in the
SSN likely contained isofunctional proteins, Cgr2 was aligned with characterized enzymes and additional selected proteins within the corresponding clusters of the SSN, and the alignment was analyzed for the presence of conserved active site residues involved in substrate binding, activation andproton transfer (Leys et al., 1999; Rohman et al., 2013; Bogachev et al., 2012; Knol et al., 2008;
Reid et al., 2000).

E. lenta and cgr2 abundance and prevalence
E. lenta and cgr2 prevalence were determined using the copy number abundance (gene copies/cell)as derived from Metaquery2 (Nayfach et al., 2015) using the median abundance from individuals
with repeated sampling. E. lenta abundance was determined from a single copy E. lenta marker
gene described elsewhere (elnmrk1) (Bisanz et al., 2018). Matches were required to have a minimum 90% nucleotide identity and query/target coverage. Reconstruction of metagenomic cgr2
sequences was carried out by quality trimming reads from 96 metagenomes with >0.001 proportional abundance of E. lenta or >1 fold coverage using default sliding window settings with Trimmomatic (Bolger et al., 2014) and extracting reads which mapped to the cgr cluster and associatedintergenic space (2957889..2968387) in the reference DSM 2243 assembly with Bowtie 2. These
were assembled and annotated as above. Alignments to Cgr2 in metagenomic coding sequenceswere filtered by a global alignment identity of >80% to position 333 ± 60 residues. For assemblyfree variant calling, reads were filtered for a minimum mapping quality of 10 and a pileup was created (SAMtools). 49 metagenomes had at least one read mapping to the variant position (2959294).
Variants were called when > 50% of reads at a site supported an alternative sequence from the reference. Conservation of nucleotide sequence in isolates was independently confirmed via Sanger
sequencing (GENEWIZ, San Francisco, CA, USA) using the following primers: cgr2_fwd (TGCAATCAAGACAACCACGA), cgr2_internal (TCGGTGTACAACCACAATGC), and cgr2_rev (GTTGCGCTGTGATTAGACTG). PCR was carried out with high-fidelity Q5 enzyme (New England Biolabs, Ipswich, MA).
To validate metagenomics inquiries, qPCR analysis with double-dye probes was carried out in a
duplexed fashion using the following primers and probes: ElentaUni_F (GTACAACATGCTCC
TTGCGG), ElentaUni_R (CGAACAGAGGATCGGGATGG), ElentaUni_Probe ([6FAM]TTCTGGC
TGCACCGTTCGCGGTCCA[BHQ1]), cgr2_F (GAGGCCGTCGATTGGATGAT), cgr2_R (ACCG
TAGGCATTGTGGTTGT), and cgr2_probe ([HEX]CGACACGGAGGCCGATGTCG[BHQ1]). Reactions
were carried out in triplicate using 10 mL reactions with 200 nM primers and probes using BioRad
Universal Probes Supermix on a BioRad CFX 384 thermocycler according to the manufacturer’s suggested settings for fast cycles with a 60 ˚C annealing temperature. The estimated assay detection
limit based on spike-in experiments is 1.4  103 GE/g after accounting for DNA extraction. Humansamples were collected for the purpose of microbiome analysis as part of the following registeredstudies: NCT03022682, NCT01967563, and NCT01105143 and approved by their respective institutional review boards. DNA was extracted with variable methods using either MoBio Power Soil

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Microbiology and Infectious Disease(QIAGEN, Germantown, MD), Qiagen Fast Stool (QIAGEN, Germantown, MD), or Promega Wizard(Promega, Madison, WI) SV 96 kits.

Statistical analysis
All statistical analysis was carried out using either Student’s t-test as implemented in Graphpad Prism
version 7 (La Jolla, CA, USA) or R version 3.4.0 using appropriate base functions for Welch’s t-test,
Pearson and Spearman correlations, and ANOVA with multcomp version 1.4–6 for Dunnett’s multiple comparison test. Graphing was carried out with Graphpad Prism and R using ggplot2 version
2.2.1. Skewedness was calculated using the R package Moments version 0.14.

Acknowledgements
We thank the National Institute of Advanced Industrial Science and Technology (Tokyo, Japan) for
providing the Rhodococcocus erythropolis L-88 strain and pTipQ expression vectors, José Miguel
Sahuquillo-Arce (Hospital Universitari i Politecnic La Fe, Spain) for providing the Eggerthella lenta
Valencia strain, and Emma Allen-Vercoe (University of Guelph, Canada) for isolating and supplying
additional strains of Eggerthella lenta. We are indebted to Patrick Bradley, Stephen Nayfach, Katherine Pollard, and Jack Nicoludis for pivotal discussions; and to Lauren Rajakovich for comments on
the manuscript. We are also indebted to Stephen Nayfach for supplying metagenomes from the
Metaquery2 set for further analysis. This work was supported by the National Institutes of Health(PJT: R01HL122593; MP: GM111978), the Searle Scholars Program (EB:12-SSP-243; PJT:SSP-2016–1352), a Fellowship for Science and Engineering from the David and Lucille Packard Foundation(EB:2013–39267), the George W. Merck Fellowship (EB:27–14), the Bill and Melinda Gates Foundation (EB:OPP1158186), and the UCSF Department of Microbiology and Immunology (PJT). PJT is a
Chan Zuckerberg Biohub investigator and a Nadia’s Gift Foundation Innovator supported, in part,by the Damon Runyon Cancer Research Foundation (DRR-42–16). PJT is also supported by the UCSF
Program for Breakthrough Biomedical Research (partially funded by the Sandler Foundation). JEB isa Natural Sciences and Engineering Research Council of Canada postdoctoral fellow. NK received
funding from the National Institutes of Health (GM095450-01), the Smith Family Graduate Scienceand Engineering Fellowship, and a National Science Foundation Graduate Research Fellowship(DGE1144152).

Additional information
Competing interests
Peter J Turnbaugh: PJT is on the scientific advisory board for Seres Therapeutics, WholeBiome, andKaleido, and has active research funding from Medimmune. In the past year PJT has consulted for
GLG and MEDAcorp. Emily P Balskus: EPB is a consultant for Merck Research Labs, Novartis, andKintai Therapeutics. The other authors declare that no competing interests exist.

Funding
FunderGrant reference number

AuthorSmith family

Graduate Science andEngineering Fellowship

Nitzan Koppel

National Science Foundation

Graduate student
fellowship, DGE1144152

Nitzan Koppel

National Institutes of HealthTraining grant, GM09545001

Nitzan Koppel

Natural Sciences and Engineering Research Council of
CanadaPostdoctoral fellowship

Jordan E Bisanz

National Institutes of Health

GM111978Maria-Eirini Pandelia

National Institutes of Health

R01HL122593

Peter Turnbaugh

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Searle Scholars Program

SSP-2016-1352; EB:12-SSP243

UCSF Department of Microbiology and Immunology
Damon Runyon Cancer Research Foundation

Peter TurnbaughPeter Turnbaugh

DRR-42-16

Chan Zuckerberg Biohub

Peter TurnbaughPeter Turnbaugh

University of California, San
Francisco

Program for BreakthroughBiomedical ResearchPeter TurnbaughDavid and Lucile Packard
Foundation2013-39267

Emily P Balskus

George W. Merck Fellowship27-14

Emily P Balskus

Bill and Melinda Gates Foundation

OPP1158186

Emily P Balskus

Searle Scholars Program

12-SSP-243

Emily P Balskus

The funders had no role in study design, data collection and interpretation, or thedecision to submit the work for publication.
Author contributions
Nitzan Koppel, Conceptualization, Formal analysis, Investigation, Methodology, Writing—originaldraft, Writing—review and editing; Jordan E Bisanz, Data curation, Formal analysis, Investigation,
Visualization, Writing—original draft, Writing—review and editing; Maria-Eirini Pandelia, Formal analysis, Investigation, Methodology, Writing—review and editing; Peter J Turnbaugh, Conceptualization, Resources, Formal analysis, Supervision, Funding acquisition, Investigation, Methodology,
Writing—original draft, Project administration, Writing—review and editing; Emily P Balskus, Conceptualization, Resources, Supervision, Funding acquisition, Writing—original draft, Project administration, Writing—review and editing
Author ORCIDs
Nitzan Koppel
http://orcid.org/0000-0002-8399-2943
Jordan E Bisanz
http://orcid.org/0000-0002-8649-1706
Maria-Eirini Pandelia
http://orcid.org/0000-0002-6750-1948
Peter J Turnbaugh
https://orcid.org/0000-0002-0888-2875
Emily P Balskus
https://orcid.org/0000-0001-5985-5714
EthicsHuman subjects: Human samples were collected for the purpose of microbiome analysis as part of
the following registered studies: NCT03022682, NCT01967563, and NCT01105143 and approvedby their respective institutional review boards.

Decision letter and Author response
Decision letter https://doi.org/10.7554/eLife.33953.043
Author response https://doi.org/10.7554/eLife.33953.044

Additional files
Supplementary files. Transparent reporting form
DOI: https://doi.org/10.7554/eLife.33953.038

Data availability
Data used was taken from http://metaquery.docpollard.org/ (job ids: 703, 805, 807, 808-813).

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