Functional Redundancy in the Hydroxycinnamate CatabolismPathways of the Salt Marsh Bacterium Sagittula stellata E-37

Ashley M. Frank,a* Michelle J. Chua,a Christopher A. Gulvik,a* Alison Buchana

aDepartment of Microbiology, University of Tennessee, Knoxville, Tennessee, USA

ABSTRACT The hydroxycinnamates (HCAs) ferulate and p-coumarate are among themost abundant constituents of lignin, and their degradation by bacteria is an essen-tial step in the remineralization of vascular plant material. Here, we investigate the ca-tabolism of these two HCAs by the marine bacterium Sagittula stellata E-37, a memberof the roseobacter lineage with lignolytic potential. Bacterial degradation of HCAs is of-ten initiated by the activity of a hydroxycinnamoyl-coenzyme A (hydroxycinnamoyl-CoA)synthase. Genome analysis of S. stellata revealed the presence of two feruloyl-CoA (fcs)synthase homologs, an unusual occurrence among characterized HCA degraders. In or-der to elucidate the role of these homologs in HCA catabolism, fcs-1 and fcs-2 were dis-rupted using insertional mutagenesis, yielding both single and double fcs mutants.Growth on p-coumarate was abolished in the fcs double mutant, whereas maximum cellyield on ferulate was only 2% of that of the wild type. Interestingly, the single mutantsdemonstrated opposing phenotypes, where the fcs-1 mutant showed impaired growth(extended lag and �60% of wild-type rate) on p-coumarate, and the fcs-2 mutantshowed impaired growth (extended lag and �20% of wild-type rate) on ferulate, point-ing to distinct but overlapping roles of the encoded fcs homologs, with fcs-1 primarilydedicated to p-coumarate utilization and fcs-2 playing a dominant role in ferulate utiliza-tion. Finally, a tripartite ATP-independent periplasmic (TRAP) family transporter wasfound to be required for growth on both HCAs. These findings provide evidence forfunctional redundancy in the degradation of HCAs in S. stellata E-37 and offer importantinsight into the genetic complexity of aromatic compound degradation in bacteria.

IMPORTANCE Hydroxycinnamates (HCAs) are essential components of lignin and areinvolved in various plant functions, including defense. In nature, microbial degrada-tion of HCAs is influential to global carbon cycling. HCA degradation pathways arealso of industrial relevance, as microbial transformation of the HCA, ferulate, cangenerate vanillin, a valuable flavoring compound. Yet, surprisingly little is known ofthe genetics underlying bacterial HCA degradation. Here, we make comparisons topreviously characterized bacterial HCA degraders and use a genetic approach tocharacterize genes involved in catabolism and uptake of HCAs in the environmen-tally relevant marine bacterium Sagittula stellata. We provide evidence of overlap-ping substrate specificity between HCA degradation pathways and uptake proteins.We conclude that S. stellata is uniquely poised to utilize HCAs found in the complexmixtures of plant-derived compounds in nature. This strategy may be commonamong marine bacteria residing in lignin-rich coastal waters and has potential rele-vance to biotechnology sectors.

KEYWORDS bacterial catabolism, ferulate, hydroxycinnamates, p-coumarate

Constituting up to 30% of the cell wall of vascular plants, lignin represents the mostabundant aromatic polymer on Earth and offers potential as a highly valuable

source of renewable carbon for numerous applications, including hydrocarbon fuel (1)and synthetic chemical production (2). Lignin is generated from the radical polymer-

Received 17 August 2018 Accepted 18September 2018

Accepted manuscript posted online 21September 2018

Citation Frank AM, Chua MJ, Gulvik CA, BuchanA. 2018. Functional redundancy in thehydroxycinnamate catabolism pathways of thesalt marsh bacterium Sagittula stellata E-37.Appl Environ Microbiol 84:e02027-18. https://doi.org/10.1128/AEM.02027-18.

Editor M. Julia Pettinari, University of BuenosAires

Copyright © 2018 American Society forMicrobiology. All Rights Reserved.

Address correspondence to Alison Buchan,abuchan@utk.edu.

* Present address: Ashley M. Frank, Departmentof Entomology, Cornell University, Ithaca, NewYork, USA; Christopher A. Gulvik, Centers forDisease Control and Prevention, Atlanta,Georgia, USA.

A.M.F. and M.J.C. contributed equally to thiswork.

PHYSIOLOGY

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ization of three hydroxycinnamyl alcohols (p-coumaryl [H], coniferyl [G], and sinapyl [S]alcohols), resulting in a structurally heterogeneous polymer harboring a variety ofaromatic substituents linked through stable carbon-carbon and ether bonds (3). De-spite this chemical complexity, many fungi and a limited number of bacteria have beendemonstrated to degrade lignin in nature (4). Primary depolymerization of lignin istypically accomplished by fungi through nonspecific oxidation driven by extracellularoxidoreductases (5–7). This initial enzymatic step liberates a pool of lower-molecular-weight aromatics that can serve as carbon and energy sources for microbes withappropriate ring fission pathways (8–10).

Among the most abundant and valuable aromatic compounds associated withlignin are the hydroxycinnamates (HCAs) p-coumarate (H unit derivative) and ferulate(G unit derivative) (11, 12). Both HCAs are essential to plant integrity and offeradditional potential for industrial and pharmaceutical applications (13). Ferulate andp-coumarate are common in crop plants, such as fruits, vegetables, and coffee (14, 15),but are most notably abundant in grasses (16). While p-coumarate is mostly esterifiedto other phenylpropanoid units of lignin (17), ferulate cross-links lignin to cell wallpolysaccharides through ether and ester bonds (18). These ferulate cross-links havebeen identified as a primary source of recalcitrance in grass lignocellulose (19). Asidefrom providing a refractory architecture, HCAs belong to a family of phenylpropanoiddefense molecules released in response to microbial invasion (12, 20). Not surprisingly,microbes with the ability to degrade HCAs are able to deflect these phenylpropanoid-derived plant defenses, leading to enhanced virulence in plants (21). A more detailedunderstanding of the genes involved in HCA degradation across different organismsmay therefore inform strategies to mitigate plant disease. In addition to supportingplant health, HCAs also harbor value from human health and industrial perspectives, asthey can serve as dietary antioxidants (22–24) and as precursors for a variety ofcommercial products. For instance, microbial strains have been engineered to convertp-coumarate into precursors for thermoplastics, flavorings, and cosmetics (25). Similarly,the bacterial degradation of ferulate, which often generates vanillin as an intermediate(26), has been exploited for the biotechnological production of commercial vanillinflavoring (27–29). Insight into the genes and pathways that modify these HCAs can thusguide technologies to generate commercially relevant products.

To date, there have been two major pathways described for the aerobic bacterialdegradation of HCAs, both of which are initiated by coenzyme A (CoA) activation of theHCA carboxylate side chain to generate a hydroxycinnamoyl-CoA (Fig. 1). In bothinstances, the CoA addition is performed by a hydroxycinnamoyl-CoA synthase orligase. The difference in the two described HCA pathways lies in the mechanismemployed for removal of the acetate moiety, with one pathway proceeding through a�-oxidative route (30, 31) and the other through a non-�-oxidative route that generatesan aldehyde intermediate (32, 33). While ferulate can be degraded through either the�-oxidative (30) or aldehyde (32–34) pathway, p-coumarate is expected to be catabo-lized solely through the aldehyde pathway (21, 32). The aldehyde pathways for ferulateand p-coumarate are homologous, differing only in the meta-ring substituent (ferulate,meta-OCH3; p-coumarate, meta-H) of the starting material and intermediates. Thus,after deacetylation in the aldehyde branch, the ferulate pathway generates vanillicaldehyde and then vanillate, whereas the p-coumarate pathway generatesp-hydroxybenzaldehyde and then p-hydroxybenzoate. All known HCA pathwaysconverge to form the central intermediate protocatechuate, which undergoes ringcleavage, with products ultimately feeding into the tricarboxylic acid (TCA) cycle (8).In the few strains in which the genetics of HCA catabolism has been elucidated, thegenes are typically colocalized and coregulated (21, 35). For example, in Agrobacteriumfabrum, all ferulate degradation genes are localized to a single catabolon (30).

For most bacteria with described HCA degradation pathways, the relevant CoA synthaseis either reported specifically for ferulate, or more generically, for hydroxycinnamates, asmany characterized CoA synthases can act upon both ferulate and p-coumarate, as well asa variety of structurally related compounds. The hydroxycinnamoyl-CoA ligase from Acin-

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FIG 1 Select HCA degradation pathways initiated by CoA addition. The left branch exhibits the �-oxidative pathwaythrough which ferulate is degraded. The right branch shows the aldehyde pathway through which both ferulate andp-coumarate can be processed. E-37 locus tags are provided at reactions for which there are strong homologs tofunctionally validated proteins and/or genome annotations. The designated proteins (NCBI RefSeq accession numbers) areas follows: SSE37_12324 (WP_005859342), SSE37_24399 (WP_005857142), SSE37_12349 (WP_005859350), SSE37_24394(WP_005857140), SSE37_12329 (WP_005859343), SSE37_12319 (WP_005859340), SSE37_18837 (WP_040603982), andSSE37_02815 (WP_005862022). Hcs, hydroxycinnamoyl-CoA synthase; Ech, enoyl-CoA hydratase; Acd, acyl-CoA dehydro-genase; Bkt, �-ketothiolase; Adh, aldehyde dehydrogenase; PobA, p-hydroxybenzoate oxygenase; VanA, vanillate-O-demethylase. HMPHP-CoA, 4-hydroxy-3-methoxyphenyl-�-hydroxypropionyl-CoA; HPHP-CoA, 4-hydroxyphenyl-�-hy-droxypropionyl-CoA; HMPKP-CoA, 4-hydroxy-3-methoxyphenyl-�-ketopropionyl-CoA. Dashed arrows indicate reactionsunique to the �-oxidative pathway, light solid lines represent reactions unique to the aldehyde pathways, and bold solidlines indicate reactions shared between the two branches. Figure is adapted from Overhage et al. (34) and Lowe et al. (21).

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etobacter baylyi ADP1 (HcaC) can transfer CoA to p-coumarate, ferulate, and caffeate (32).Similarly, the annotated feruloyl-CoA ligase (FerA) from Sphingobium sp. strain SYK-6 hasbeen demonstrated to convert p-coumarate, ferulate, caffeate, and sinapate (33). Althoughdirect evidence is not yet available, the CoA ligase (Atu1416) from A. fabrum is also likelyto accept both ferulate and p-coumarate based on transcriptomics data (36) andgrowth profiles from deletion constructs (30).

The marine bacterium Sagittula stellata E-37 has been developed as a model for thestudy of HCA degradation and the catabolism of other plant-derived aromatic com-pounds in coastal ocean environments (37–39), where the dissolved organic carbonpool is highly aromatic in nature (40). This isolate has been shown to selectively attachto and partially mineralize a synthetic form of lignin and demonstrates robust growthon a variety of aromatic monomers, including ferulate and p-coumarate (38, 39, 41).However, the pathways utilized by this strain to degrade these HCAs are unknown. Asdescribed herein, the genome of S. stellata harbors two open reading frames (ORFs)with significant homology to hydroxycinnamoyl-CoA synthases (both annotated asferuloyl-CoA synthases [Fcs]). Since Fcs proteins have been shown to have broadsubstrate specificities across para-substituted HCAs and are encoded by a single genein other bacteria (33, 42), the presence of two disparately localized fcs homologs in S.stellata is an intriguing phenomenon and suggests that dedicated pathways forp-coumarate and ferulate exist in this strain. This genetic arrangement in S. stellatapresents insight into the complexity of HCA catabolism in lignin-rich coastal marineenvironments.

RESULTSS. stellata possesses two disparately located hydroxycinnamoyl-CoA synthase

homologs. The S. stellata genome was queried for the presence of hydroxycinnamoyl-CoA synthase homologs through alignments to functionally validated protein se-quences from Acinetobacter baylyi ADP1 (GenBank accession no. AAL54850), Agrobac-terium fabrum C58 (NCBI RefSeq accession no. NP_354423), and Sphingobium sp. SYK-6(GenBank accession no. BAK67177). BLASTp alignments of the selected proteinsagainst the S. stellata genome identified two ORF products, here referred to as Fcs-1(WP_005859342) and Fcs-2 (WP_005857142), which showed highest sequence similar-ity to A. baylyi and A. fabrum Fcs proteins. Fcs-1 showed moderately higher sequenceidentity and expectation (E) values than both of these previously characterized Fcsproteins (identities, 55 and 40%; E values � 0 and e�147 for A. fabrum and A. baylyi,respectively) relative to Fcs-2 (identity, 40 and 36%; E values � e�140 and e�124 for A.fabrum and A. baylyi, respectively). No significant homology was evident between thevalidated Fcs from Sphingobium and either of the S. stellata Fcs sequences. S. stellataFcs-1 and Fcs-2 sequences share 41% identity and 57% similarity to one another andpossess conserved sequence motifs diagnostic of hydroxycinnamoyl synthases (i.e.,acyl-activating enzyme [AAE] consensus, putative AMP binding site, putative active site,and putative CoA binding site motifs).

S. stellata ORFs predicted to be involved in downstream conversions for the�-oxidative branch for HCA degradation were also identified (Fig. 2 and Table 1).BLASTp alignment of the �-oxidative acyl-CoA dehydrogenase (Atu1415) from A.fabrum (30) against the S. stellata genome yielded homology to SSE37_24394 (identity,51%; E value � e�79), annotated as a 3-hydroxy-2-methylbutyryl-CoA dehydrogenase,which lies just upstream of fcs-2. Interestingly, another acyl-CoA dehydrogenase(Atu1414) exists within the A. fabrum HCA catabolon but was shown not to be essentialto ferulate degradation in that system (36). This protein demonstrates homology to theS. stellata acyl-CoA dehydrogenase SE37_12329 (identity, 55%; E value � e�138) locatednext to fcs-1. Last, the A. fabrum HMPKP �-ketothiolase, Atu1421, strongly aligns to S.stellata SSE37_12319 (identity, 82%; E value � e�180), annotated as a 4-hydroxyphenyl-beta-ketoacyl-CoA hydrolase, located adjacent to fcs-1.

The two fcs genes and several of the genes potentially involved in downstreamconversions reside in two distinct genetic loci on the S. stellata chromosome (Fig. 2). S.

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stellata harbors ORFs with high sequence similarity to many of the gene productswithin the A. fabrum catabolon (Table 1), nearly all of which are within 5 kb of fcs-1. Thesingle exception is an acyl-CoA dehydrogenase, which is adjacent to fcs-2. Interestingly,fcs-2 is immediately upstream of an operon that encodes the recently characterizedbenzoyl-CoA oxidation (Box) pathway, through which benzoate is degraded by thisstrain (S. stellata E-37) (39). Additionally, fcs-2 is just downstream from a three-genecassette anticipated to encode a tripartite ATP-independent periplasmic (TRAP) familytransporter, which consists of a substrate-binding protein (DctP; SSE37_24379) and twomembrane-bound proteins (DctQM; SSE37_24384 and SSE37_24389) (43) (Fig. 2). Theproximity of the TRAP gene cassette to these catabolic genes suggests that theencoded transporter may facilitate the uptake of aromatic compounds.

Though S. stellata can use vanillin as a sole carbon source (see Fig. S3 in thesupplemental material), genomic evidence for the aldehyde pathway, which generatesvanillin as an intermediate, is not strong in S. stellata. Homology searches to charac-terized vanillin dehydrogenases (i.e., LigV [GenBank accession no. BAK65381] fromSphingobium sp. SKY-6 and HcaB [CAG68567] from A. baylyi) result in moderate-to-weaksimilarities (maximum identities, LigV � 37%, E value � e�76; HcaB � 34%, E value

FIG 2 Genetic landscape of the two fcs loci in S. stellata. Locus tags for predicated HCA-related degradation genes in E-37 and associated A. fabrum homologs(denoted with Atu prefix) are provided with percent protein identity above the corresponding gene arrows. Dark-pink arrows represent genes involved in the�-oxidative and/or the aldehyde pathway described in Fig. 1. Light-pink arrows indicate genes involved only in the �-oxidative pathway. bkt, �-ketothiolase;fcs, feruloyl-CoA synthase; acd, acyl-CoA dehydrogenase; ech, enoyl-CoA hydratase. Other functionally related genes are coordinately color coded and includea predicted operon for cobalamin biosynthesis (SSE37_12299, SSE37_12304, and SSE37_12309); the dct operon (SSE37_24379, SSE37_24384, and SE37_24839),predicted to encode a C4-dicarboxylate TRAP transporter; and a portion of the box pathway (SSE37_24404, SSE37_24409, SSE37_24414, SSE37_24419, andSSE37_24424) encoding enzymes for benzoate degradation. Dark gray represents genes for which a defined role in either HCA degradation or another pathwayis not predicted with the following numbers: 1 indicates SSE37_12314, a protein of unknown function (DUF 3237); 2 indicates SSE37_12334, annotated as analdo/ketoreductase; 3 indicates SSE37_12339, annotated as a flavin adenine dinucleotide (FAD)-dependent oxidoreductase; and 4 indicates SSE37_12344,annotated as a MarR-like transcriptional regulator, as discussed in the text. The complete box operon is provided in Fig. S6.

TABLE 1 S. stellata homologs to A. fabrum HCA catabolism proteins

A. fabrum locus(NCBI RefSeqaccession no.)

S. stellata homolog locus(NCBI RefSeq accession no.) Putative function Coverage (%)

Amino acididentity (%) E value

Atu1414 (NP_354421) SSE37_12329 (WP_005859343) Acyl-CoA dehydrogenase 83 54 3e�138

Atu1415 (NP_354422) SSE37_24394 (WP_005857140) Acyl-CoA dehydrogenase 100 51 1e�79

Atu1416 (NP_354423) SSE37_12324 (WP_005859342) Feruloyl-CoA synthase 98 55 0.0Atu1416 (NP_354423) SSE37_24399 (WP_005857142) Feruloyl-CoA synthase 97 40 2e�140

Atu1417 (NP_354425) SSE37_12349 (WP_005859350) Enoyl-CoA hydratase 96 67 6e�126

Atu1421 (NP_354429) SSE37_12319 (WP_005859340) �-Ketothiolase 100 82 8e�180

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�e�82) to a number of putative aldehyde dehydrogenases encoded in the S. stellatagenome, none of which are adjacent to either fcs locus.

Genetic analysis reveals overlapping but distinct roles of fcs genes in HCAdegradation. To assess the contributions of the putative fcs homologs to hydroxycin-namate catabolism in S. stellata, mutant strains lacking either or both functional copiesof fcs-1 and fcs-2 were generated, and the phenotypes on a suite of substrates weredetermined. The three mutants (fcs-1, fcs-2, and fcs-1 fcs-2) displayed wild-type growth onthe control nonaromatic and aromatic carbon sources, acetate and p-hydroxybenzoate(POB), respectively (Fig. S4A and B). In S. stellata, POB is converted to the central interme-diate protocatechuate (PCA), which then undergoes ring fission (39). Hence, wild-typegrowth on POB is a strong indicator that the ring cleavage pathway through which all otheraromatic acids tested here are funneled (37–39) is unaffected by the mutations.

Growth curve analyses on ferulate revealed that while the fcs-1 mutant demon-strated wild-type growth, the fcs-2 mutant showed an extended lag phase (Fig. 3A) andsignificantly depressed growth rate (22% of wild-type rate; P � 0.01; Fig. S5). Yet, theculture ultimately reached wild-type cell densities in stationary phase (Fig. 3A). The fcs-1fcs-2 double mutant elicited a severely depressed growth phenotype on this substrate,with a maximum absorbance reading only �30% of that of the wild-type cultures. Asferulate imparts a maroon coloration to the medium that may inflate absorbancereadings, viable counts were also performed as an additional measure of cell growth.Maximum viable cell abundances of the double mutant grown on ferulate were only2% of those of the wild-type cultures (1.81 � 107 [� 1.56 � 106] CFU/ml versus 7.27 �

108 [� 7.64 � 107] CFU/ml, respectively). Consistent with this finding, only 30% of theprovided ferulate was taken up by the double mutant. In contrast to the doublemutant, ferulate concentrations were drawn below the limit of detection for all otherstrains (Fig. 3C).

The individual fcs mutants showed opposing phenotypes on p-coumarate. The fcs-2mutant displayed wild-type growth, while the fcs-1 mutant strain showed a lag phasedouble that of wild-type cultures (Fig. 3B) and a significantly decreased exponential-growth rate (61% of wild-type rate; P � 0.01; Fig. S5). Both single mutants ultimatelyreached wild-type cell densities on this substrate, whereas the double mutant failed to

FIG 3 Catabolism of hydroxycinnamates by S. stellata E-37 and the fcs mutants. Growth of strains on ferulate (A) or p-coumarate (B), withcorresponding concentrations of ferulate (C) or p-coumarate (D) in cell-free spent medium. Wild type is shown with black closed circles, the fcs-1mutant with blue open circles, the fcs-2 mutant with red open circles, and the fcs-1 fcs-2 double mutant with yellow open circles. Averages froma minimum of three biological replicates are shown, along with one standard deviation from the mean.

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grow (Fig. 3B). The abolished growth of the double mutant on p-coumarate wascorroborated by the lack of p-coumarate depletion from the medium (Fig. 3D). For allstrains and substrates, the extracellular concentrations of substrates were consistentwith the growth assays (Fig. 3).

Given that many aromatic acids are toxic to bacterial cells (44, 45), a parallel set ofgrowth experiments were performed using a Biolog array, which included a tetrazoliumdye that is responsive to cellular respiration-associated redox changes (46). Here, cellswere incubated in 96-well plates supplemented with ferulate, p-coumarate, or controlcompounds (this study), in the presence of a proprietary tetrazolium-based dye, andmonitored for colorimetric changes indicative of dye reduction upon cellular respira-tion. The Biolog assays were conducted to cross-validate the optical density (OD)-basedgrowth studies, as measurement of respiration provides a supplemental analysis of cellgrowth. The general trends from these assays are consistent with absorbance-basedmeasures of growth, with the exception of the ferulate Biolog measurements, whichmay be due to interference of the maroon color imparted by ferulate (Fig. S4).Regardless, the viable count and HCA depletion assays agree that the double mutantcannot effectively oxidize ferulate, while the fcs-2 mutant is moderately delayed in itsprocessing.

Gene expression assays indicate that fcs genes are growth substrate inducible.To complement the mutational analysis, reverse transcription-quantitative PCR (RT-qPCR) was performed to assess fcs gene expression when S. stellata was grown onferulate or p-coumarate. Both the fcs-1 and fcs-2 genes were significantly upregulatedduring growth on both HCAs relative to growth on the nonaromatic substrate acetate(P � 0.005). However, the relative gene expression of fcs-1 far exceeded that of fcs-2.The fcs-1 expression levels from p-coumarate- and ferulate-grown cells were 10-foldhigher than those of fcs-2 during growth on either HCA (Fig. 4).

To expand our understanding of fcs gene regulation in S. stellata, additional se-quence analysis was performed to identify putative transcriptional regulators. A puta-tive MarR-type regulator (NCBI RefSeq accession no. WP_005859349, SSE37_12344) ispresent within the fcs-1 region (Fig. 2). Alignment of the S. stellata MarR to this proteinyields very weak identity (35%). Along those same lines, there is no notable similaritybetween the S. stellata predicted regulatory protein and the predicted regulator of theA. fabrum hca catabolon, GntR (NCBI RefSeq accession no. NP_354412, Atu1405). Here,the S. stellata putative protein (NCBI RefSeq accession no. WP_005861787) with highestsequence similarity to GntR demonstrates low identity (30%) and poor E value (e�6).Furthermore, it is not located near either fcs gene. A single putative regulator(SSE37_24449) is found within the fcs-2 region. This gene is located within and

FIG 4 fcs gene expression for S. stellata grown on two hydroxycinnamates. Gene expression wasrelativized to growth on 7 mM acetate and normalized to three housekeeping genes. Blue bars representfcs-1 expression, and red bars represent fcs-2 expression. Identical letters above bars indicate nonsignif-icant values, whereas nonidentical letters represent a significant difference (*, P � 0.005). Averages froma minimum of three biological and two technical replicates are shown, along with one standard deviationfrom the mean.

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cotranscribed with the box operon but is not anticipated to be cotranscribed with fcs-2given the intergenic distance (�500 bp) (Fig. S6).

Identification of a transporter for hydroxycinnamates in S. stellata. ThoughFcs-1 appears to play a more dominant role in p-coumarate degradation and Fcs-2 inferulate degradation, our results suggested overlapping specificities, implying compen-satory roles for each protein when provided a nonpreferred HCA substrate. We nextsought to determine whether this apparent relaxation of specificity was extended toproteins involved in the uptake of these compounds.

We focused our efforts on the dctPQM cassette located adjacent to the fcs-2 gene(Fig. 2). Initial RT-qPCR studies of ferulate- and p-coumarate-grown cells showed dctPgene expression to be elevated 13-fold (�2-fold) and 23-fold (�14-fold), respectively,relative to acetate-grown cultures. To functionally determine the role of this putativearomatic transporter, dctP (SSE37_24379) was interrupted using insertional mutagen-esis. The dctP mutant was unable to grow on either ferulate or p-coumarate (Fig. S7),supporting its indispensable role in HCA transport. Partial restoration of activity wasevident with a wild-type dctP-complemented strain (Fig. S7).

DISCUSSION

Despite the abundance of hydroxycinnamates (HCAs) in lignin-rich environmentsand the demonstrated importance of bacteria in the recycling of this material, relativelylittle is known of the bacterial catabolic pathways used to remineralize these com-pounds. Using the model marine heterotrophic bacterium Sagittula stellata E-37, weprovide compelling evidence for the presence of multiple HCA degradation pathwayswith overlapping substrate specificities. We argue that this genetic arrangement allowsthe strain to be poised to utilize these compounds in nature, where they are found incomplex mixtures of aromatics.

Our data support a primary role for fcs-1 in p-coumarate catabolism and fcs-2 in ferulatecatabolism in S. stellata, with evidence of overlapping functionalities between the twoenzymes encoded by these genes. The ability of each single mutant to ultimately reachwild-type cell densities on nonoptimal substrates suggests a compensatory function foreach of their gene products. The concept of overlapping substrate specificity for this classof enzymes is not novel, as many aromatic CoA ligases have been demonstrated totransform a variety of structurally related compounds. As previously mentioned, thehydroxycinnamoyl-CoA ligases from A. baylyi (HcaA) (32), Sphingobium sp. SYK-6 (FerA) (33),and A. fabrum (Atu1416) (30, 36) demonstrate broad substrate specificity. Although thesewell-documented CoA ligases convert multiple substrates, these strains do not possessmultiple hydroxycinnamoyl-CoA ligase homologs, nor is there evidence for compensatoryfunction. Accordingly, single fcs gene knockouts were sufficient to prevent HCA utilizationin both A. fabrum and Sphingobium spp. (30, 33).

The expanded substrate range of the two Fcs proteins from S. stellata is furthercorroborated by transcriptional analyses of the wild-type strain, which show that bothfcs-1 and fcs-2 are upregulated in cultures grown on either HCA. However, transcrip-tional responses to growth on HCAs were different, perhaps in part due to differencesin their basal (noninduced) expression levels. S. stellata grown with acetate as the solecarbon source had 5.2� (�0.9�) more fcs-2 transcripts than fcs-1 transcripts. Thisdifference in basal gene expression suggests that these genes are under differenttranscriptional regulation. Consequently, S. stellata appears to be better poised torapidly oxidize ferulate with Fcs-2. If substantiated, these findings could indicate that S.stellata is primed to respond to ferulate as a growth substrate.

The difference in expression of fcs-1 and fcs-2 in S. stellata suggests distinctregulation of the two genes. This is perhaps intuitive given the physical separation ofthe two fcs genes within the S. stellata chromosome. There are annotated regulatorswithin both fcs genomic regions that could potentially serve this function; however,convincing genomic evidence is lacking. In A. baylyi, a MarR-type regulator (GenBankaccession no. AAP78949) is responsible for regulating an hca operon (32). While aputative MarR-type regulator (SSE37_12344) is located in the S. stellata fcs-1 region, it

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shares little sequence similarity with the A. baylyi characterized protein. A singleputative regulator (SSE37_24449) is found within the fcs-2 region but is not predictedto regulate HCA degradation based on gene expression assays. This ORF (SSE37_24449)is predicted to encode a transcriptional regulator for benzoate catabolism and has beenaccordingly designated BoxR. SSE37_24449 shares 44% (E value � e�68) sequenceidentity with two transcriptional regulators of benzoate catabolism in Azoarcus sp.strain CIB (BoxR [Ga0098266_114713] and BzdR [Ga0098266_111630]) (47, 48), thebacterium for which the novel aerobic box pathway was first described (49). Finally,SSE37_24449 contains the Walker-A motif [consensus sequence, GLRGAGK(T/S)], whichhas been proposed to specifically interact with benzoyl-CoA (47). Thus, there is cur-rently no evidence to support the involvement of SSE37_24449 or SSE37_12344 in thedirect genetic regulation of HCA catabolism in S. stellata.

The apparent relaxed substrate specificity for the HCA catabolic proteins appears toextend to proteins involved in transport. The TRAP transporter gene dctP (SSE37_24379) isrequired for growth on both ferulate and p-coumarate. The upregulation of dctP in S.stellata cultures grown on p-coumarate or ferulate provides additional evidence that theproducts of the dctPQM cassette act on both p-coumarate and ferulate. The broad speci-ficity of TRAP family members is well documented and includes aromatic acids (43). InComamonas sp. strain DJ-12, the transport of 4-chlorobenzoate and related compounds ismediated by a TRAP family transporter (50). The Rhodopseudomonas palustris TRAPsubstrate-binding protein TarP has been shown to bind not only ferulate and p-coumaratebut also caffeate and cinnamate (51). No other aromatic acid transporters have yet beenidentified in S. stellata. and very few genes with sequence similarity to transporter proteinshave been identified in aromatic compound catabolism gene clusters in this strain (39).

The ability of the fcs double mutant to sustain limited growth on ferulate isintriguing. It is conceivable that S. stellata is able to access the methoxy side chain offerulate, which is absent in p-coumarate, to support restricted growth. The spectro-photometric assay indicates the double mutant is capable of taking up �30% of theprovided substrate, though cellular growth is not consistent with complete utilizationof this amount of ferulate. This could occur through removal of the entire methoxygroup to generate p-coumarate, an unusable product for the double mutant, orthrough removal of the methyl group to yield caffeate (as observed in plants [52]), asubstrate S. stellata is unable to catabolize (39). Although documentation of methoxyutilization from lignin-derived aromatics is typically observed with anaerobic methano-gens or acetogens (53, 54), S. stellata harbors a putative vanillate-O-demethylase(SSE37_02815) that could presumably remove the methyl group from ferulate. Theenergy and biomass derived from a C1 methyl group would be considerably less thanthat of C10 ferulate, which is consistent with the growth phenotype of the mutant.Furthermore, although results presented here indicate that S. stellata predominantlydegrades ferulate via a �-oxidative mechanism, it is feasible that additional parallelnot-yet-characterized pathway(s) for ferulate degradation exist in this strain and couldexplain this restricted growth phenotype.

While this study suggests a dominant role of fcs-1 for p-coumarate utilization andfcs-2 for ferulate utilization, an important ecological finding is the ability of both ofthese genes to be upregulated by its noncognate HCA substrate. The cross-substrateutilization may reflect the diverse substrate pools to which the strain is adapted. Theeastern U.S. coastal salt marshes from which the strain was isolated (55) are rich indecaying plant matter, principally derived from the cordgrass Spartina alterniflora,which is rich in ferulate and p-coumarate (56). Indeed, plant detritus is the principlesource of dissolved organic carbon in these territories (57). These coastal marshes arealso dominated by members of the Roseobacter lineage, to which S. stellata belongs(58). The ability to degrade aromatic compounds is common among lineage members,as is the presence of multiple (�6) ring-cleaving pathways (59). Roseobacters, ingeneral, and S. stellata, in particular, appear to be uniquely adapted to accessinglignin-derived carbon. S. stellata is able to mineralize synthetic lignin (38) and isadapted to growth on mixtures of aromatic compounds. The strain demonstrates a

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synergistic growth response when provided mixtures of two lignin degradation prod-ucts, benzoate and p-hydroxybenzoate (39). This phenomenon has been shown forother roseobacters (39) and indicates a positive relationship between substrate com-plexity and aromatic compound utilization in this group of bacteria. In this context, themultisubstrate specificity of the two hydroxycinnamoyl-CoA synthases encoded by S.stellata seems conducive to its lifestyle, as it appears to have evolved an arsenal of toolsto degrade the wide breadth of aromatics in its surroundings.

These insights into HCA catabolism of the marine bacterium S. stellata unveil themodularity and complexity of approaches for aromatic compound degradation withinthis organism. Through a continued understanding of the reactions used to transformlignin-derived compounds, there is potential for uncovering novel physiologies, path-ways, and enzymes that could be of industrial use, as many valuable products can begenerated from lignin (2) and its HCA derivatives (25–27). The ease of handling and theexpanding genetic tractability of S. stellata and other roseobacters (60) offer an excitingopportunity for future engineering applications.

MATERIALS AND METHODSStrains and growth conditions. Sagittula stellata E-37 was previously isolated from pulp mill effluent

and enriched on Indulin AT, a commercially available Kraft lignin generated from pulp mills, as describedby Gonzalez et al. (38). Unless otherwise noted, wild-type S. stellata and any mutant derivatives wereroutinely maintained at 30°C in YTSS medium (per liter, 2.5 g yeast extract, 4 g tryptone, 15 g sea salts[Sigma-Aldrich, St. Louis, MO]). Escherichia coli strains used for cloning and conjugation experiments weremaintained at 37°C in Luria-Bertani broth (per liter, 10 g tryptone, 5 g yeast extract, 10 g NaCl). Antibioticswere added to the E. coli growth medium to maintain selective pressure at 50 �g/ml kanamycin or 10�g/ml tetracycline. S. stellata growth assays were routinely performed in marine basal medium (MBM)containing 1.5% (wt/vol) Sigma sea salts, 225 nM K2HPO4, 13.35 �M NH4Cl, 71 mM Tris-HCl (pH 7.5), 68�M Fe-EDTA, trace metals and vitamins (38), and carbon (2 mM aromatic and 10 mM acetate, unlessotherwise noted). Strains were incubated at 30°C, in the dark, with shaking. All glassware was combustedin an ashing furnace (type F62700; Barnstead Thermolyne) for 4 to 24 h prior to use to eliminate tracecarbon. Seeding densities were �1 � 106 CFU/ml, and no-carbon-added controls were run in parallel forcomparison. Growth was monitored using a spectrophotometer at 540 nm. All strains used in this studyare listed in Table 2.

Generation of S. stellata mutants. A S. stellata fcs-1 (SSE37_12324) mutant strain was producedusing marker exchange mutagenesis following modifications of previously described procedures (61–63).Briefly, a 402-bp region upstream and a 585-bp region downstream of fcs-1 was amplified to generate“A” and “B” amplicons, respectively. During PCR amplification, a 21-bp scar sequence was added throughthe internal primers, generating complementary appendages for overlap extension PCR. PCR mixtures foreach A and B reaction included 1� GoTaq buffer (Promega, Madison, WI), 200 �M deoxynucleotidetriphosphates (Promega), 6 �M internal primer, 0.6 �M external primer, 0.025 U/�l GoTaq DNA poly-merase (Promega), and 1 to 2 ng/�l S. stellata genomic DNA. Thermocycling conditions consisted of a3-min denaturation at 95°C, followed by 31 cycles of 30 s at 95°C, 30 s at 56°C, 30 s at 72°C, and a finalextension for 5 min at 72°C. The A and B amplicons were subsequently used as the template DNA duringoverlap extension PCR to generate a fused “AB” product. This PCR was performed with 1� FailSafe bufferF (Epicentre, Madison, WI), 6 �M external primer A, 6 �M external primer B, 0.05 U/�l FailSafe enzymemix (Epicentre), 1 to 2 ng/�l amplicon A, and 1 to 2 ng/�l amplicon B. The thermocycling conditions werethe same as used previously, except with a 10-min final extension. The resulting product was cloned intopCR2.1-TOPO (Invitrogen, Carlsbad, CA). Positive clones were selected on LB-kanamycin (50 �g/ml) andverified by sequence analysis. The AB insert was then subcloned into the mobilizable suicide plasmidpARO180 (ATCC 77123), through shared HindIII and XbaI sites, generating plasmid pARO180_12324, andtransformed into chemically competent E. coli JM109. Positive clones were selected on LB-ampicillin (50�g/ml) agar and verified through amplification and sequencing. The sacB-kanR cassette from pRMJ1 (63) wasinserted into the middle of the AB insert at the SalI site, generating plasmid pARO180_12324_SK. Thecomplete A-sacB-kanR-B vector construct was then transformed into the mating strain, E. coli S17-1. Thesuicide vector was delivered to S. stellata through biparental mating with E. coli S17-1(pARO180_12324_SK)on a YTSS agar plate. S. stellata transconjugants were enriched from the mating mixture by requiring growthon 5 mM p-hydroxybenzoate (a restrictive substrate for E. coli) and 50 �g/ml kanamycin. Allelicreplacement was confirmed by PCR amplification across the chromosomal region of interest usingprimers 12324_AsacF/12324_BkmR (Table 3) and sequencing of the PCR product. PCR for this amplifi-cation was performed using the LongRange PCR kit (Qiagen, Valencia, CA), according to the manufac-turer’s instructions, with the following thermocycling conditions: initial denaturation at 93°C for 3 min,followed by 30 cycles of 93°C denaturation for 15 s, 57°C annealing for 30 s, 68°C elongation for 12 min,and no final extension.

The fcs-2 (SSE37_24399) mutant was generated using insertional mutagenesis with the pKNOCK-Kmplasmid (64). A 231-bp internal region of SSE37_24399 (bp positions 983 to 1213) was cloned intopCR2.1-TOPO, subcloned into pKNOCK-Km to generate plasmid pKNOCK-Km_24399, and then intro-duced into the E. coli mating strain BW20767 (65). Conjugation of BW20767(pKNOCK-Km_24399) with

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wild-type S. stellata was performed as described above. Single-crossover homologous recombinationresulted in integration of the entire plasmid into the fcs-2 gene. The fcs-1 fcs-2 double mutant wasgenerated by cloning the same 231-bp fragment of fcs-2 into pKNOCK-Tc (64) (making pKNOCK-Tc_24399), and biparental mating of the E. coli BW20767 harboring this plasmid with the fcs-1 mutantstrain of S. stellata. Different plasmids were used for chromosomal integration of fcs-1(pARO180) andfcs-2(pKNOCK) to eliminate the chance of crossover between homologous sections of the plasmidsduring generation of the double knockout.

A dctP (SSE37_24379) mutant was generated using insertional mutagenesis with the pKNOCK-Kmvector harboring a 292-bp internal fragment (positions 423 to 714 bp) of dctP and following proceduresoutlined above. To confirm the role of SSE37_24379, complementation of the dctP with the broad-host-range plasmid pRK415 (66) harboring the wild-type dctP gene was attempted. The phenotypes ondifferent substrates were determined by growth assays on HCAs and their intermediates.

All chromosomal disruptions were confirmed by PCR amplification and sequencing of junction sites.All plasmids and primers used in generation of mutants are listed in Tables 2 and 3, respectively.

Biolog phenotypic microarrays of fcs mutants. Custom phenotypic microarrays were performed toassay for redox activity of strains when grown on defined carbon sources (10 mM acetate and 2 mM eacharomatic monomer [p-hydroxybenzoate, ferulate, and p-coumarate]). All fcs mutants and wild-type S.stellata strains were precultured in MBM with 10 mM acetate and supplemented with kanamycin andtetracycline, as necessary. Assays were performed in 96-well flat-bottom plates (Costar) containing 1�MBM, 1� Biolog dye mix G (Hayward, CA), 107 cells/ml S. stellata, and a single carbon source. Plates wereincubated at 30°C in an OmniLog reader (Biolog), where digital colorimetric readings were capturedevery 30 min to monitor the intensity of dye reduction over time, recorded in OmniLog units/time.

RNA isolation and reverse transcription. For total RNA extraction, strains were grown on 7 mMacetate or 2 mM HCA. Total RNA was extracted using the RNeasy minikit (Qiagen), with slight modifi-

TABLE 2 Plasmids and strains used in this study

Plasmid or strain Descriptiona Source or reference

PlasmidspCR2.1-TOPO TA plasmid for cloning InvitrogenpRMJ1 Plasmid harboring sacB-kanR cassette for marker exchange mutagenesis 63pARO180 Mobilizable suicide plasmid for marker exchange mutagenesis D. Park (ATCC 77123)pARO180_12343 pARO180 containing fused flanking regions of fcs-1 (SSE37_12324) cloned into

XbaI and HindIII sites of the MCSThis study

pARO180_12324_SK pARO180_12324 with sacB-kanR from pRMJ1 inserted into the middle of the fcs-1insert through designed SalI site

This study

pKNOCK-Km Mobilizable suicide plasmid harboring kanamycin resistance gene; used for site-directed mutagenesis via chromosomal integration

64 (Addgene 46262)

pKNOCK-Km_24399 pKNOCK-Km with small internal region (231 bp) of fcs-2 cloned into BamHI andXhoI sites of the MCS

This study

pKNOCK-Tc Mobilizable suicide plasmid harboring tetracycline resistance gene; used for site-directed mutagenesis via chromosomal integration

64 (Addgene 46259)

pKNOCK-Tc_24399 pKNOCK-Tc with small internal region (231 bp) of fcs-2 cloned into the BamHIand XhoI sites of the MCS

This study

pKNOCK-Km_24379 pKNOCK-Km with 292-bp internal region of dctP cloned into the HindIII andBamHI sites of the MCS

This study

pRK415 Broad-host-range plasmid for DNA cloning in Gram-negative bacteria used fordctP complementation

66

pRK415_24379 pRK415 containing a 1.7-kb HindIII-BamHI DNA fragment that includes the region579 bp upstream of dctP start codon and 83 bp downstream of dctP stopcodon

This study

StrainsE-37 Wild-type Sagittula stellata E-37 58E-37 fcs-1::sacB-kanR E-37 fcs-1 mutant where fcs-1 (SE37_12324) is replaced with sacB-kanR from

pARO180_12324_SKThis study

E-37 fcs-2::pKNOCK-Km E-37 fcs-2 mutant where fcs-2 (SSE37_24399) is interrupted by pKNOCK-Km_24399 This studyE-37 fcs-1::sacB-kanR fcs-2::pKNOCK-Tc E-37 fcs-1 fcs-2 double mutant generated by interrupting fcs-2 (SSE37_24399) with

pKNOCK-Tc_24399 in strain E-37 fcs-1::sacB-kanRThis study

E-37 dctP::pKNOCK-Km E-37 dctP mutant wherein dctP (SSE37_24379) was interrupted bypKNOCK-Km_24379

This study

E-37 dctP-pKNOCK-Km_dctP::pRK415 E-37 dctP mutant complemented with pRK415_24379 This studyE-37 dctP-pKNOCK-Km_pRK415 E-37 dctP mutant transformed with empty pRK415 vector This studyE. coli TOP10 Cloning strain for pCR2.1-TOPO plasmid InvitrogenE. coli JM109 Subcloning and transformation strain InvitrogenE. coli S17-1 Mating strain for marker exchange mutagenesis ATCC 47055E. coli BW20767 Mating strain for pKNOCK mutagenesis, derived from S17-1 65 (ATCC 47084)

aMCS, multiple cloning site.

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TABLE 3 Primers used in this study

Primer name Sequencea Functionb

12324_A_ex GATCACTGCGCCGATCTT Primer pair amplifies A region (402 bp upstream of12324) with 21-bp scar SalI site appended to the3= end; A_ex primer is also used with B_ex forcrossover PCR to generate the A-scar-B product

12324_A_in gattcgaggagcgatagagctGTCGACTTCTCCTCCTGGCGTTTTAG12324_B_ex GGAGCCCAACAGCATCAT Primer pair amplifies B region (585 bp downstream

of 12324) with 21-bp scar appended to the 5=end; B_ex primer is also used with A_ex forcrossover PCR to generate the A-scar-B product

12324_A_in agctctatcgctcctcgaatcGACCGGCGCCCGGTT12324_Asac_F GGATCAGGATGTCGTCGTTC Primer pair binds E-37 DNA upstream and

downstream of fcs-1; used to verify replacementof gene with sacB-kanR cassette

12324_Bkm_R CAGCTTCTCCTCGATTCGCT24399_int_F TGGTGTTTTATGCAGGCGCG Primer pair amplifies 231-bp internal region of

fcs-2 (SSE37_24399) used for cloning intopKNOCK plasmids

24399_int_R TTTCGCAGCGCATGTCTTCGpKNOCKKm_752_F ACGGCTGACATGGGAATTCC Primer pair binds around MCS of pKNOCK-Km to

amplify inserts off plasmidpKNOCKKm_901_R GCGGAATTAATTCGACGCGTCpKNOCKTc_425_F GAATTCCCCTCCACCGCGG Primer pair binds around MCS of pKNOCK-Tc to

amplify inserts off plasmidpKNOCKTc_558_R TGATCAAGCTGACGCGTCCT24399_884_F GAACGCTGGCCTTCAACGTG Primer pair binds E-37 DNA upstream and

downstream of the 231-bp internal region offcs-2; used to verify insertion of pKNOCKplasmids in fcs-2 of E-37

24399_1383_R GAAATCCTCCGAAATCCGCCC24379_423_F TCTGGAAGGCATGAAGATCC Primer pair amplifies 292-bp region of dctP

(SSE37_24379) used for cloning into pKNOCK-Km24379_714_R ATCGATCACCTCCTGCAGAT24379_367966_F TCTAGAAGCTTTGTTTGATGGATCAACGCACT Primer pair amplifies 1.7-kb HindIII-BamHI DNA

fragment which encompasses the region 579 bpupstream of dctP start codon and 83 bpdownstream of dctP stop codon

24379_369669_R TGCGTAGGATCCATCAGGATCAGGAACGTCAGC24379_690_F AGAGGATCTGCAGGAGGTGA Primer pair amplifies 188-bp region of dctP

(SSE37_24379) used for quantifying dctPexpression in WT E-37 on hydroxycinnamic acids

24379_877_R ACGTCTCGTAGATCGGGTTG12324_562_F CTGACAGCGACACACCTGAC Amplifies 181-bp region of (SSE37_12324) for

quantifying fcs-1 expression in WT E-37 and fcs-2mutant

12324_742_R CCTTCAGGAAGGTGAAGCAC24399_1249_F CTCAACGACCCGAAGAAGAC Amplifies 187-bp region of fcs-2 (SSE37_24399) for

quantifying fcs-2 expression in WT E-37 and fcs-1mutant

24399_1435_R ACAATTCCAGACGCAGGTTC15096_340_F GCCCATATCTGGTTCCTCAA Primer pair amplifies 159-bp region of rpoC

(SSE37_15096) for use as housekeeping gene 1of 3 in RT-qPCR

15096_498_R TTCCTCTTCGGTCAGCATCTmap_13553_620_F GCATGTTCTTCACCATCGAG Primer pair amplifies 166-bp region of map

(SSE37_13553) for use as housekeeping gene 2of 3 in RT-qPCR

map_13553_785_R GCGGGAGAGAGGGTAAAGATalaS_ 05000 _97_F GATCCGACGCTTATGTTCGT Primer pair amplifies 151-bp region of alaS

(SSE37_05000) for use as housekeeping gene 3of 3 in RT-qPCR

alaS_ 05000_247_R TGTAACCGACGTTGTCCAGAaThe 21-bp scar sequences are in lowercase. The SalI site is underlined.bWT, wild type.

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cation; cells were lysed by vortexing at maximum speed for 10 min with 0.2 g of low-binding 200-�mzirconium beads (OPS Diagnostics, LLC, Lebanon, NJ). Genomic DNA was removed using the TurboDNA-free kit (Ambion, Austin, TX), as per the manufacturer’s instruction. cDNA synthesis was performedwith Moloney murine leukemia virus (MMLV) reverse transcriptase (Invitrogen) using 500 ng total RNA,as described in the product manual.

Gene expression analyses. RT-qPCR was performed using 1� SYBR Premix Ex Taq (Perfect Real Time)(TaKaRa Bio, Inc., Otsu, Japan) in 25-�l volumes consisting of 12.5 �l of 1� SYBR premix, 10 �l cDNA, and 1.25�l forward and 1.25 �l reverse primers (10 �M). Genes of interest were normalized to three previouslyvalidated housekeeping genes, rpoC, map, and alaS (39). Quantitative PCR (qPCR) amplification included a95°C denaturation for 3 min, followed by 40 cycles of 95°C for 20 s, 57°C for 20 s, and 72°C extension for 15s, and a final extension at 72°C for 5 min. Melt curves were obtained between 50 and 100°C at 1°C per s withreadings every 1°C. cDNA was diluted with 10 mM Tris (pH 8.0), and endpoint PCR was performed prior toRT-qPCR to ensure that the threshold cycle (CT) values for each gene and replicate were between 15 and 30cycles. Efficient genomic DNA removal (CT values �5 cycles) was confirmed by including no-RT controls.Technical and biological replicates were included in triplicate for each gene. RT-qPCR analyses were per-formed using qBase (67) to normalize and relativize gene transcripts.

Analysis of substrate concentrations. The concentrations of p-coumarate and ferulate weremonitored in cell-free supernatants from S. stellata cultures spectrophotometrically. The wavelengths atwhich each substrate absorbed maximally in minimal medium were empirically determined by perform-ing wavelength scans from 200 to 700 nm, with 0.002% sample (1 ml total volume, diluted with sterileMilli-Q water). Molar extinction coefficients for each substrate in the minimal medium were calculated tobe 0.21 liters · mol�1 · cm�1 for ferulate and 0.40 liters · mol�1 · cm�1 for p-coumarate. To quantifyextracellular concentrations of substrates, culture aliquots were passaged through a 0.22-�m filter, andabsorbances at 283 nm (p-coumarate) and 308 nm (ferulate) were monitored with a Beckman DU 800UV-Vis spectrophotometer. Concentrations were calculated from 10-point standard curves (0.3 to 3.0mM), with r2 values of 0.97 for ferulate and 0.99 for p-coumarate (Fig. S1). Full (200 to 700 nm)wavelength scans were performed for all cultures. The shape of the absorbance curves providesadditional evidence of the specificity of the approach (Fig. S2).

Genome analyses. Protein sequences from organisms with experimental validation of function wereused in homology searches against the S. stellata genome. These searches were performed using theBLASTp algorithm at NCBI.

SUPPLEMENTAL MATERIAL

Supplemental material for this article may be found at https://doi.org/10.1128/AEM.02027-18.

SUPPLEMENTAL FILE 1, PDF file, 5.3 MB.

ACKNOWLEDGMENTSWe are grateful to Elizabeth Fozo (UTK) for providing guidance on transcriptional

analysis of the box operon. This research was supported by a grant from the NationalScience Foundation (OCE-1357242). Additionally, A.M.F. and C.A.G. were supported aspart of the Center for Direct Catalytic Conversion of Biomass to Biofuels (C3Bio), anEnergy Frontier Research Center funded by the U.S. Department of Energy, Office ofScience, Office of Basic Energy Sciences, award number DE-SC0000997.

A.M.F., C.A.G., and A.B. developed initial project objectives and experimental design.A.M.F. generated the fcs mutants and M.J.C. constructed the dctP mutant. A.M.F. andM.J.C. performed mutant growth characterizations. M.J.C. performed RT-qPCR assays,with statistical support from C.A.G. A.M.F. led fcs data analysis and manuscript prepa-ration, with support from M.J.C. and A.B.

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