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Journal of Bacteriology logoLink to Journal of Bacteriology
. 2014 Jul;196(14):2670–2680. doi: 10.1128/JB.01556-14

Tolerance of a Phage Element by Streptococcus pneumoniae Leads to a Fitness Defect during Colonization

Hilary K DeBardeleben a, Elena S Lysenko a, Ankur B Dalia c, Jeffrey N Weiser a,b,✉
PMCID: PMC4097588  PMID: 24816604

Abstract

The pathogenesis of the disease caused by Streptococcus pneumoniae begins with colonization of the upper respiratory tract. Temperate phages have been identified in the genomes of up to 70% of clinical isolates. How these phages affect the bacterial host during colonization is unknown. Here, we examined a clinical isolate that carries a novel prophage element, designated Spn1, which was detected in both integrated and episomal forms. Surprisingly, both lytic and lysogenic Spn1 genes were expressed under routine growth conditions. Using a mouse model of asymptomatic colonization, we demonstrate that the Spn1− strain outcompeted the Spn1+ strain >70-fold. To determine if Spn1 causes a fitness defect through a trans-acting factor, we constructed an Spn1+ mutant that does not become an episome or express phage genes. This mutant competed equally with the Spn1− strain, indicating that expression of phage genes or phage lytic activity is required to confer this fitness defect. In vitro, we demonstrate that the presence of Spn1 correlated with a defect in LytA-mediated autolysis. Furthermore, the Spn1+ strain displayed increased chain length and resistance to lysis by penicillin compared to the Spn− strain, indicating that Spn1 alters the cell wall physiology of its host strain. We posit that these changes in cell wall physiology allow for tolerance of phage gene products and are responsible for the relative defect of the Spn1+ strain during colonization. This study provides new insight into how bacteria and prophages interact and affect bacterial fitness in vivo.

INTRODUCTION

Streptococcus pneumoniae, or the pneumococcus, is a Gram-positive opportunistic pathogen responsible for ∼1 million deaths worldwide each year (http://www.who.int/immunization/monitoring_surveillance/burden/estimates/Pneumo_hib/en/). Although it is a leading source of infection, the pneumococcus is most commonly found in a commensal relationship with its human host (1). Pneumococcal colonization is characterized by sequential and overlapping episodes that each last for days to months. Colonizing pneumococci provide the reservoir of organisms causing disease and are the main source of bacteria for host-to-host transmission (2). Therefore, colonizing pneumococci are likely to be the major focus of selective pressure and adaptation for the organism. During colonization, pneumococci encounter various challenges, one of which is predation by bacteriophages. Up to 70% of clinical isolates of S. pneumoniae harbor genetic elements resembling prophages based on the presence of the phage-encoded lysin, which is a homologue of the chromosomally encoded cell wall amidase, LytA (3). How these prophage elements affect the physiology of their pneumococcal host during colonization has not been fully explored.

The predator-prey dynamic between phages and the bacteria they infect has been an important factor driving bacterial evolution (4). This arms race has resulted in the evolution of phages that promote the fitness of their bacterial host, including its survival within a mammalian host and ability to cause disease (5). For example, prophages may encode toxins or other virulence factors that increase the ability of their bacterial host to persist during infection. Phages may also impact genetic diversity within a species through lysogenic conversion or horizontal gene transfer, which facilitates bacterial adaptation to the host environment (e.g., antibiotic resistance) (6).

In contrast to the positive effects of phages on bacterial pathogenesis, few studies have assessed the negative effect of phages on bacterial survival during infection. Aspects of the temperate phage life cycle that could have a negative impact are phage lysis, increased burden of DNA and protein synthesis, and cis-acting effects on genes flanking the phage attachment site. The variety of phage resistance mechanisms employed by bacteria indicates that negative effects of carrying prophage are a significant factor in bacterial fitness (7).

In the case of the pneumococcus, prophage SV-1 has been shown to enhance biofilm formation by increasing amounts of extracellular DNA (8). Similarly, pneumococcal prophage MM1 has been shown to increase bacterial adherence to human epithelial cells in culture (9). However, the contribution of these or other pneumococcal phages to positive or negative effects on bacterial fitness in vivo has not been demonstrated.

Using a newly identified prophage, Spn1, found in a highly successful pneumococcal lineage (10), and a well-established mouse model (11), we examined how the prophage affects S. pneumoniae during colonization. The bacterial isolate harboring Spn1 was obtained from a human challenge study and shown to be fit for colonizing the natural host for up to 122 days (10) and the murine nasopharynx for 21 days (11). We find that the Spn1 element is stably present and transcriptionally active in this isolate. However, the presence of Spn1 correlates with a negative impact on bacterial fitness during colonization. We also provide evidence that Spn1 is associated with changes in the cell wall physiology of its host. We posit that this change is a mechanism for tolerance of phage products, which may be the underlying cause of this fitness defect.

MATERIALS AND METHODS

Bacterial growth conditions.

Unless noted otherwise, bacteria were grown in C+Y medium, pH ∼6.8 (12), without shaking in sealed tubes at 37°C. Growth was monitored via absorbance readings at 620 nm. Bacteria were plated on blood agar plates or tryptic soy medium (TS) plus catalase (4,741 U/plate) (Worthington Chemicals). C+Y was supplemented with streptomycin (200 μg/ml), kanamycin (500 μg/ml), erythromycin (1 μg/ml), neomycin (5 μg/ml), or spectinomycin (200 μg/ml) where indicated.

Sequencing of Spn1.

P1121, a bacterial isolate from a human challenge study (Table 1), was sequenced on an Illumina HiSeq and de novo assembled using CLC's Main Workbench (CLC bio). Two assembled contigs containing prophage coding regions were identified by NCBI's BLAST program. The gap between the two contigs was sequenced using Sanger sequencing technology. Once the sequence was complete, the genome was submitted to RAST for annotation (13).

TABLE 1.

Bacterial strains

Strain no. Designation Background Descriptiona Source or reference
P1121 Spn1+ Human challenge study isolate, serotype 23F 11
P1397 Spn1+ Smr P1121 Smr spontaneous mutation This study
P2197 MM1− TIGR4 10
P2198 MM1+ TIGR4 10
P2282 Spn1+ ΔlytA P1121 lytA::Spr This study
P2308 Spn1+ Δmml P1121 mml::Kmr This study
P2345 Spn1+ ΔSP_1563 P1121 SP_1563::Spr This study
P2348 Spn1+ ΔSP_1564 P1121 SP_1564::Emr This study
P2352 Spn1− Smr P1397 Unmarked in-frame Spn1 deletion, Smr This study
P2373 Spn1+ ΔpblB P1121 pblB::Emr This study
P2379 Spn1− ΔlytA P2385 lytA::Spr transformed with P2282 This study
P2380 Spn1+ Δmml ΔlytA P2308 mml::Kmr lytA::Spr transformed with P2282 This study
P2385 Spn1− P2352 Transformed with P1121 to remove Smr This study
P2386 Spn1+ Δint Δmml P2385 mml::Kmr int::Emr transformed with P2387 This study
P2387 Spn1+ Δint Δmml P2308 mml::Kmr int::Emr transformed with P2388 This study
P2388 Spn1+ Δint P1121 int::Emr This study
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a

Smr, streptomycin resistant; Spr, spectinomycin resistant; Kmr, kanamycin resistant; Emr, erythromycin resistant.

RNA isolation and qRT-PCR.

RNA was isolated using the TRIzol reagent and protocol (Life Technologies), with modifications. Bacteria were harvested at mid-log phase, and a 2:1 volume of RNAprotect bacterial reagent (Qiagen) was added to the culture. Bacteria were then resuspended in TRIzol, frozen, and then disrupted by bead beating for 5 min. RNA was isolated by phenol-chloroform extraction as written in the TRIzol protocol. After resuspension of the RNA in RNase-free water, samples were treated with DNase (Qiagen) for 30 min at 37°C. DNase was inactivated at 65°C for 10 min before RNA was used to make cDNA. Quantitative reverse transcription-PCR (qRT-PCR) was carried out using a high-capacity cDNA reverse transcription kit (Applied Biosystems). Quantitative PCR was performed with the Power SYBR green PCR master mix (Applied Biosystems). mRNA levels were quantified using ΔCT (threshold cycle) values and compared to an internal control, gyrA. Primers are listed in Table 2.

TABLE 2.

Primers for mutant construction and qRT-PCR

Primer Gene Direction Sequence Source
AttF SP_1563 Forward TGTGGGTGGTGGTCCTGTCG This study
AttR SP_1564 Reverse ATGCCAAACTGGCCCGTCAC This study
EGP4 int Forward GAAGATAGGAGGATAAACTGG 22
EGP9 mml Reverse AACTGCAGAAATTGTTCTTTCACCGCAGG 22
Spn1-2 mml Reverse CATTATCCATTAAAAATCAAACAACTGCAGAAATTGTTCTTTCACCGCAGG This study
Spn1-3 Janus Forward CCTGCGGTGAAAGAACAATTTCTGCAGTTGTTTGATTTTTAATGGATAATG This study
Spn1-4 Janus Reverse CCAGTTTATCCTCCTATCTTCCTTTCCTTATGCTTTTGGAC This study
Spn1-5 int Forward GTCCAAAAGCATAAGGAAAGGAAGATAGGAGGATAAACTGG This study
1563-1 SP_1561 Forward TATAGCCGCCCGGTGTCTGG This study
1563-2 SP_1563 Reverse CACTTTATTAATTTGTTCGTATGTATTCAGTTTCTCCTTTGTTTTTTCTAGTCAGTTTAT This study
1563-3 SpecR Forward ATAAACTGACTAGAAAAAACAAAGGAGAAACTGAATACATACGAACAAATTAATAAAGTG This study
1563-4 SpecR Reverse CGAATCCTATGTGACTCGTGGTTCTTTTTTCCCGAGCTCGAATTGACGCGGATCC This study
1563-5 SP_1563 Forward GGATCCGCGTCAATTCGAGCTCGGGAAAAAAGAACCACGAGTCACATAGGATTCG This study
1563-6 mml Reverse CGCTGCAACAGGCTGGCAGA This study
1564-1 int Forward TGGAGAAGAAAACTCCCCAGGCA This study
1564-2 SP_1564 Reverse GGTGCAAGTCAGCACGAACGATATAAAGCAACCCCTTGAATTATCAA This study
1564-3 ermR Forward ATAATTCAAGGGGTTGCTTTATATCGTTCGTGCTGACTTGCACC This study
1564-4 ermR Reverse GATGCGCATTATACAGGTGAAAAATGAGTAACGTGTAACTTTCCAAAT This study
1564-5 SP_1564 Forward ATTTGGAAAGTTACACGTTACTCATTTTTTCACCTGTATAATGCGCATC This study
1564-6 SP_1565 Reverse GGCCAGCCGTCGAGTAGTGC This study
mml-1 SP_1563 Forward GCAGCCTTTTATGCCCACCTACGCC This study
mml-2 mml Reverse CTCTGGAATAGGCATAGACACTATCCACCGCAGGCTCAGGCTTGCGG This study
mml-3 kanR Forward CCGCAAGCCTGAGCCCTGCGGTGGATAGTGTCTATGCCTATTCCAGAG This study
mml-4 kanR Reverse GACGCATGGAAAGGACGATAGGGACACGTTTTTGTGGTGAGAAAC This study
mml-5 mml Forward GTTTCTTCACCACAAAAACGTGTCCCTATCGTCCTTTCCATGCGTC This study
mml-6 pblB Reverse CTGAGCATAAGGAATAGGAGGTGTG This study
int-1 int Forward ATGTGGATGGAAGAACTTTCCAAC This study
int-2 int Reverse GGTGCAAGTCAGCACGAAACAAGTGCTTTTATTTCTTGCATGGT This study
int-3 ermR Forward ACCATGCAAGAAATAAAAGCACTTGTTTCGTGCTGACTTGCACC This study
int-4 ermR Reverse GTACTTTTGGTGATATTCTCGACGATTAAGAGTAACGTGTAACTTTCCAAAT This study
int-5 int Forward ATTTGGAAAGTTACACGTTACTCTTAATCGTCGAGAATATCACCAAAAGTAC This study
int-6 int Reverse AGTATCTAATTTATTGACCAGTTTCTCCTCC This study
PblB-1 ORF61 Forward GAAGCTGGTAAAGAGGCGGT This study
PblB-2 pblB Reverse ATGGTGCAAGTCAGCACGAATCCTTGGCAAACATGGCTCT This study
PblB-3 ermR Forward AGAGCCATGTTTGCCAAGGATTCGTGCTGACTTGCACCAT This study
PblB-4 ermR Reverse ATGGTTTCAGCCTCTTGAGCTAGTAACGTGTAACTTTCCAA This study
PblB-5 pblB Forward TTGGAAAGTTACACGTTACTAGCTCAAGAGGCTGAAACCAT This study
PblB-6 ORF55 Reverse GTCAGGATTGCCCTCTGCAT This study
LytA-1 dinF Forward TTGGCTAGTTCGACAGATGGTTAC This study
LytA-2 lytA Reverse CACTTTATTAATTTGTTCGTATGTATTCTTCTACTCCTTATCAATTAAAACAACTCA This study
LytA-3 specR Forward TGAGTTGTTTTAATTGATAAGGAGTAGAAGAATACATACGAACAAATTAATAAAGTG This study
LytA-4 specR Reverse ATGCGCTGTTCTGATTTGAAAGACATTCCCCCGAGCTCGAATTGACGCGGATCC This study
LytA-5 lytA Forward GGATCCGCGTCAATTCGAGCTCGGGGGAATGTCTTTCAAATCAGAACAGCGCAT This study
LytA-6 SP_1935 Reverse TGAGTTCTATTGGCATTTTCTCTG This study
RTmmlF mml Forward CCACTCAACAGGCAACCGTA This study
RTmmlR mml Reverse CTTCCGTTGTTCACAGGACC This study
RTholinF holin Forward TTGCGTAAAGGCGGAGAGAA This study
RTholinR holin Reverse AGCTACTTGCTCAACGGCAT This study
RTPblBF pblB Forward TATCAAGGCCGAATCGGTGG This study
RTPblBR pblB Reverse GGCACCATTCCCCATACCAA This study
RTORF42F ORF42 Forward GGACGCAGACTACAGCAAGT This study
RTORF42R ORF42 Reverse CATATCGGCAGGCACGTACT This study
RTORF20F ORF20 Forward AAGAGTCAAATCGGTGGCGT This study
RTORF20R ORF20 Reverse GCTCCTTTGTCAGCAATCGAC This study
RTC1RepF C1Rep Forward TCGCACAGATAATCCTACCATCGCT This study
RTC1RepR C1Rep Reverse TGAAGCGCATAGCAGTAGAGGCA This study
RTintF int Forward CGACTTAGCGAACGTTTCCAGCCA This study
RTintR int Reverse ACGTGGGGGTCGTGGTGTCC This study
RT1563F SP_1563 Forward CGACCTCAACCGTCACAAGA This study
RT1563R SP_1563 Reverse TAGCAGCAGTCACCGATAGC This study
RT1564F SP_1564 Forward AGGTCGTTACGGGTTTCTCG This study
RT1564R SP_1564 Reverse ACCTATCTCCAGCGAGCAGA This study
RTgyrAF gyrA Forward GCCCTTTGGCAGTCCGACCA This study
RTgyrAR gyrA Reverse ACGTGGGGGTCGTGGTGTCC This study
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DNase protection assay.

Supernatants from cultures left untreated or treated with 5 μg/ml mitomycin C (Sigma-Aldrich) were filtered through a 0.22-μm-syringe filter. Samples were divided and either left untreated or treated with DNase (final concentration, roughly 270 Kunitz units/ml) (Qiagen) and incubated for 30 min at 37°C. DNase was then inactivated at 65°C for 10 min. DNA was quantified via quantitative PCR using the Power SYBR green PCR master mix (Applied Biosystems).

Generation of mutants.

Bacterial mutants were created in the P1121 background by first amplifying regions upstream and downstream of the gene of interest via PCR. For each mutant, a set of primers 1 to 6 was designed. Primers amplifying the flanking regions are designated 1 and 2 (upstream) and 5 and 6 (downstream). Antibiotic cassettes were amplified using primers 3 and 4. PCR was carried out using standard techniques with either Taq DNA polymerase (Invitrogen) and related reagents for non-cloning-related PCR assays or Platinum Pfx polymerase (Invitrogen) and related reagents for PCR related to cloning. Primers used are listed in Table 2. Primers 2 to 5 were tagged with an overlapping sequence. A DNA fragment was then created using overlap-extension PCR (14) with the amplified regions flanking the antibiotic resistance cassette of choice. Antibiotic resistance markers used were erythromycin resistance, Emr (pCR2.1-TOPO plasmid with Emr insertion from pMU1328 [15]), kanamycin resistance, Knr (Janus cassette [16]), and spectinomycin resistance, Spr (pCR2.1-TOPO with Spr insertion from pBI143 [aad9; GenBank accession number U30830] [17]). Each of the markers used contains its own promoters. This PCR fragment was transformed directly into bacteria using a transformation protocol as previously described (18). Mutants with an unmarked deletion were made via the Janus cassette as previously described (16). Primers and antibiotic cassettes used for each mutant strain are listed in Table 2. Constructs were confirmed by sequencing across junctions.

Mitomycin C assays.

Bacteria were grown to mid-log phase and then diluted to an optical density at 620 nm (OD620) of 0.1 in C+Y medium with or without 0.5 μg/ml of mitomycin C (Sigma-Aldrich). Growth at 37°C without shaking was recorded as the OD620 more than 4 h after treatment. Samples were collected for RNA isolation 2 h after addition of mitomycin C. Control strains TIGR4 (MM1−) and TIGR4M (MM1+) were obtained from V. Fischetti, Rockefeller University (Table 1).

Mouse model of colonization.

Bacteria were grown to mid-log phase (OD620 of 0.5) in C+Y medium. One ml of culture was pelleted and resuspended in 100 μl of phosphate-buffered saline (PBS). For competitive assays, bacteria were combined in a 1:1 ratio after resuspension. Ten μl of resuspended bacteria was dropped onto the nostrils of nonanesthetized wild-type female C57BL/6 mice (6 to 8 weeks of age). Inoculum counts were ∼1 × 106 CFU/ml. For competitive assays, at least 50 colonies of the inoculum were plated to determine streptomycin sensitivity or resistance and, thereby, the input ratio. Seven days after inoculation, nasal lavages were taken as previously described (19) and plated on TS with neomycin (5 μg/ml) to determine CFU counts. For competitive assays, at least 50 colonies from each mouse were tested for streptomycin resistance or sensitivity to determine an output ratio. Competitive indices (CIs) were calculated as (output ratio)/(input ratio). Statistics were calculated on log values of all competitive indices.

In vitro competitive growth assay.

Bacterial strains were grown to an OD620 of 0.5 in C+Y and then diluted to an OD620 of 0.1. One ml of each strain and 2 ml of fresh culture, for a total of 4 ml, were combined to start the assay. Cultures were incubated, without shaking, at 37°C and plated for bacterial counts at 0 h and 5 h on selective media for an input and output ratio, respectively, to calculate the competitive index.

Growth and autolysis assays.

Bacteria were diluted from mid-log-phase culture to a low starting OD620 (∼0.05), and then they were grown for 24 h in a 96-well plate in an incubating plate reader set at 37°C in 180 μl C+Y medium with 10 μl of catalase (30,000 U/ml) (Worthington Chemicals). Penicillin sensitivity was determined across a range of concentrations (50 ng/ml exceeded the MIC for both strains, defined as total inhibition of growth) under these conditions. Absorbance at 600 nm was measured every 15 min with 5 s of shaking on a low setting before every reading. To maximize autolysis, cocultures were carried out in brain heart infusion medium (BHI), and 50 μl of each of the strains was grown to mid-log phase and diluted to a low OD620 (∼0.05).

Western blotting.

Whole-cell lysates were created by resuspending bacteria (harvested at mid-log phase) in loading dye (30 mM Tris-HCl, pH 6.8, 2% SDS, 10% glycerol, and 2% β-mercaptoethanol) and incubating them at 95°C for 5 min. Lysates were loaded onto a 10% glycine gel (Mini-Protean TGX gel; Bio-Rad) and run under standard conditions. Protein was transferred to a polyvinylidene difluoride (PVDF) membrane via a semidry transfer. After transfer, the membrane was allowed to dry, blocked with 2% bovine serum albumin (BSA) solution in phosphate-buffered saline, and incubated with a 1:100 dilution of anti-LytA rabbit IgG (obtained from R. Lopez, Biological Research Center, Madrid, Spain) or a 1:40 dilution of anti-pneumolysin mouse IgG (Novacastra Laboratories Ltd., Buffalo Grove, IL) for 2 h. This was followed by incubation with an appropriate secondary antibody conjugated to alkaline phosphatase (anti-rabbit IgG or anti-mouse IgG, respectively, diluted 1:5,000 [Sigma]). Proteins were visualized with a 5-bromo-4-chloro-3′-indolylphosphate and nitroblue tetrazolium reaction.

Microscopy.

Bacteria were grown to mid-log phase in C+Y medium. Samples were blinded before images at ×400 magnification were taken on a Nikon E600 bright-field microscope. ImageJ analysis was performed using the “threshold” function followed by the “analyze particles” function to quantify particle size. For transmission electron microscopy, samples were grown to mid-log phase and then pelleted. Pelleted bacteria were high-pressure frozen, freeze substituted into acetone with 2% OsO4 and 0.1% uranyl acetate, embedded in EPON resin, cut into 70-nm sections, and imaged on a JEOL 1010 microscope on an AMT camera.

Nucleotide sequence accession number.

The sequence for the phage element Spn1 was deposited in GenBank under accession number KJ417497.

RESULTS

Identification of phage element Spn1.

Phage element Spn1 (GenBank accession no. KJ417497) was first identified in a human colonization isolate of Streptococcus pneumoniae via PCR using the prophage typing system described by Romero et al. (20). Subsequent sequencing of the entire locus revealed a novel phage element, of ∼42 kb, most closely related to phage 040922 (GenBank accession no. FR671406.1). However, the integrase (int) and lysin (mml) of Spn1 shared 99% and 95% identity, respectively, with the previously characterized pneumococcal phage MM1 (GenBank accession no. AJ302074.2). The entirety of Spn1 was not identified in any previously published pneumococcal genome; therefore, it is not likely to be common. The sequence of phage element Spn1 was annotated using RAST, which predicted 63 open reading frames. Similar to lambda phage, the genome was organized into lytic (lysis, structural, and replication genes) and lysogenic functional clusters (Fig. 1A). The Spn1 phage element integrates between two convergently transcribed genes, annotated as SP_1563 (a pyridine-nucleotide-disulfide-oxidoreductase) and SP_1564 (conserved hypothetical protein) in the TIGR4 genome (GenBank accession no. AE005672.3). The attachment site is likely to be the same attachment site for MM1, a 15-bp sequence located at the 3′ end of the coding sequence for SP_1564 (Fig. 1B) (21). Neither flanking gene's coding region is interrupted by integration of Spn1. Both flanking genes are highly conserved among S. pneumoniae genomes.

FIG 1.

FIG 1

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Genetic map and activity of Spn1. (A) The strain P1121, including the genetic element Spn1, was sequenced on an Illumina Hi-Seq and annotated using RAST. Sixty-three open reading frames were identified and are indicated as block arrows showing the direction of transcription. Genes were arranged in lytic or lysogenic clusters by putative function: lysis genes (black), structural genes (dark gray), replication genes (white), and lysogeny genes (light gray). (B) Schematic of Spn1 (circle) integration between genes SP_1563 and SP_1564 (dashed arrows show the direction of transcription). The attachment site, 15 bp at the 3′ end of the coding region of SP_1564, is indicated by a black bar. The light gray bar indicates a noncoding intergenic region. Small arrows represent the location of primers. (C) PCR products showing the presence of a circularized element (primers a and b), integration at the attachment site (primers a and d), or the attachment site lacking Spn1 (primers c and d) in the indicated strains. Size markers are indicated in base pairs. (D) Levels of mRNA relative to the housekeeping gene gyrA as determined by qRT-PCR for genes across Spn1. The position of specific genes is designated in panel A and shaded to match the functional cluster in the genetic map. Data are shown as means ± standard errors of the means (SEM).

Spn1 activity.

We designed PCR primers to amplify across the attachment site of both the prophage and the bacteria to determine if Spn1 is capable of excising (Table 2). Both sets of primers amplified a product in noninduced genomic DNA samples, indicating a background level of spontaneous excision and circularization of Spn1 from the P1121 genome (Fig. 1C). Using qRT-PCR on bacterial RNA isolated from noninduced log-phase cultures, we determined that Spn1 expresses its lysogenic genes as expected but also expresses lytic genes, including those encoding its lysin (Mml) and holins (Fig. 1D). Despite the expression of lytic genes, we were unable to identify plaques on any indicator strain using a plaque assay that was previously described for S. pneumoniae (22). Additionally, there was no evidence of phage particles using a DNase protection assay or in transmission electron micrographs of strain P1121. A selectable marker was inserted within Spn1 to determine whether the prophage could be cured, as indicated by the loss of the marker during passage. No Spn1− revertants were identified during in vitro growth (spontaneous cure rate of <0.002 per generation). Spn1 was then removed from strain P1121 using Janus cassette technology to generate an unmarked, in-frame deletion, Spn1− (16).

Spn1 induction by mitomycin C.

The P1121 lysogen was treated with the DNA cross-linking agent mitomycin C (5 μg/ml) to trigger the SOS response and activate the Spn1 prophage. Mitomycin C treatment of strain TIGR4 with or without phage MM1 served as controls. There was inhibition of growth correlating with the presence of phage MM1 but no effect of mitomycin C on the growth of strain P1121 carrying Spn1 compared to the isogenic Spn1− construct (Fig. 2A). To confirm that the mitomycin C treatment was activating the Spn1 prophage under these conditions, phage gene expression was assessed. Gene expression was greatly increased in the presence of mitomycin C for lytic and lysogenic genes across Spn1 (Fig. 2B). Thus, Spn1 is transcriptionally activated by mitomycin C, but this treatment is not sufficient to induce lysis of its bacterial host.

FIG 2.

FIG 2

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Activation of Spn1 by mitomycin C. (A) Growth of strain P1121 with or without Spn1 and strain TIGR4 with or without MM1 in C+Y medium. Mitomycin C (5 μg/ml) was added at time zero. (B) Relative transcript abundances as determined by qRT-PCR for the Spn1 gene indicated relative to the internal control, gyrA, is shown relative to treatment without mitomycin C. Data are shown as means ± SEM. One-sample t test was used to determine if the relative increase in gene expression was greater than 1. *, P < 0.05.

Spn1 causes a defect in fitness in vivo.

To determine the effect of Spn1 on its bacterial host in vivo, we used a murine nasopharyngeal colonization model. When tested individually, the Spn1+ and Spn1− strains each colonized mice at similar densities at 7 days postinoculation (Fig. 3A). This time point was chosen because it is the extent of a period of stable colonization before clearance becomes prominent (23). To compare their relative fitnesses, we competed Spn1+ and Spn1− strains 1:1 in the same model of colonization. At 7 days postinoculation, the Spn1− strain significantly outcompeted the Spn1+ strain (CI, >70) (Fig. 3B). Competitive growth in vitro did not correlate with competitive effect in vivo, indicating that Spn1 confers a fitness disadvantage that is specific to the in vivo environment (Fig. 3B). In order to differentiate strains postcolonization, one strain was marked with an rpsL mutation conferring resistance to streptomycin. Data shown is a combination of experiments with either the Spn1+ or Spn1− strain having been marked to ensure that the streptomycin resistance phenotype was not the cause of the fitness effect in vitro or in vivo.

FIG 3.

FIG 3

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Spn1 in colonization. (A) Density of Spn1+ and Spn1− in nasal lavages obtained 7 days after inoculation. The dashed line indicates the limit of detection. Comparisons between strains were made using the Mann-Whitney test. (B) Competitive indexes (CI; output ratio/input ratio) of Spn1−/Spn1+ following growth for 5 h in C+Y medium (in vitro) or 7 days of colonization (in vivo). (C) Competitive indexes for Spn1− ΔlytA/Spn1+ ΔlytA and Spn1− ΔlytA/Spn1+ ΔlytAΔmml to determine the competitive advantage without autolysis and with equivalent chain length, respectively. Each symbol is from a single mouse or culture. The CI is based on patching ≥50 colonies on selective medium to distinguish strains. One-sample t test was used to determine if CIs were significantly different from a hypothetical value of 1, which would indicate no competitive advantage or disadvantage. *, P < 0.05; ***, P < 0.001; ns, nonsignificant.

A trans-acting factor from Spn1 affects in vivo fitness.

To assess whether the effect of Spn1 on in vivo fitness was caused by cis-acting effects on the flanking genes, we generated mutants in SP_1563 and SP_1564 in strain P1121 and tested them in the colonization model. The loss of SP_1563 resulted in a defect in colonization (Fig. 4A). To determine if Spn1 was affecting transcription of the flanking genes, we compared mRNA levels of Spn1+ and Spn1− strains using qRT-PCR. Expression of SP_1563 and SP_1564 was unaffected by the presence of Spn1 (Fig. 4B). Therefore, an effect of Spn1 on either flanking gene was unlikely to be the cause of the fitness defect in vivo.

FIG 4.

FIG 4

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Spn1 gene expression is required for the fitness defect in vivo. (A) Density of Spn1+(WT), Spn1+ ΔSP_1563, and Spn1+ ΔSP_1564 in nasal lavages obtained 7 days after inoculation. Comparisons between mutant and WT strains were made by Mann-Whitney test. (B) Levels of mRNA relative to the internal control, gyrA, as determined by qRT-PCR of genes SP_1563 and SP_1564 in the presence (black bars) or absence (white bars) of Spn1. Data are shown as means ± SEM. (C) Levels of mRNA compared to the internal control, gyrA, as determined by qRT-PCR for sample phage genes in either the Spn1+ strain or the Spn1+ Δint Δmml strain. Expression was undetected (ND) in the Spn1+ Δint Δmml strain. Data are shown as means ± SEM. (D) Competitive indexes of Spn1−/Spn1+ Δint Δmml following growth for 5 h in C+Y medium or 7 days of colonization. (E) Competitive indexes of Spn1+ ΔpblB/Spn1+, Spn1−/Spn1+ Δmml, and Spn1−/Spn1+ Δint following 7 days of colonization. One-sample t test was used to determine significant differences from a hypothetical value of 1, which would indicate no competitive advantage. *, P < 0.05; **, P < 0.01; ns, nonsignificant.

We attempted to reinsert the Spn1 prophage in the Spn1− strain but were unable to identify transformants acquiring this large genetic element. By placing selectable markers within both the int and mml genes at opposite poles of the prophage, it was possible to obtain transformants of the Spn1− strain in which the entire locus was restored. However, interruption of both the int and mml genes resulted in a strain in which Spn1 no longer excised spontaneously (Fig. 1C) or transcribed genes across the entire prophage (Fig. 4C). Theoretically, this mutant should maintain any cis-acting effects on the flanking genes but lack trans-acting effects due to phage gene expression. The competitive index of the Spn1+ Δint Δmml strain versus the Spn1− strain was not significantly different from 1, indicating that the two strains compete equally (Fig. 4D). This result suggests that a trans-acting factor requiring phage gene expression is required for the effect of Spn1 on in vivo fitness. This result also eliminated the possibility that the burden of carrying an additional ∼42 kb of DNA was responsible for our observations on fitness during colonization.

Three candidate genes were identified as potential trans-acting factors in the fitness effect of Spn1. First, the phage lysin, Mml, was interrupted to create an Spn1+ Δmml strain. The competitive index of Spn1− versus Spn1+ Δmml was significantly greater than 1, indicating that the Spn1-mediated fitness effect was not due to Mml (Fig. 4E). The second candidate gene, PblB, a large phage tail protein, was removed to create the Spn1+ ΔpblB strain. Competition between Spn1+ and Spn1+ ΔpblB resulted in a competitive index not significantly different from 1, indicating no competitive advantage for either strain (Fig. 4E). Lastly, the integrase gene (int), a DNA binding protein that mediates integration of the phage (24), was interrupted to create Spn1+ Δint. The competitive index of Spn1− versus Spn1+ Δint is significantly greater than 1, indicating that Spn1− has the fitness advantage. This leads us to conclude that Mml, PblB, or integrase alone does not confer the fitness defect to the Spn1+ strain in vivo.

The presence of Spn1 correlates with delay in autolysis.

The phage lysin, Mml, and the major autolysin of S. pneumoniae, LytA, are homologous and thought to be functionally redundant cell wall amidases (22). In order to determine if autolysis is affected by the presence of Spn1, we measured bacterial growth in nutrient broth medium over a period of 24 h. Autolysis was delayed in the Spn1+ strain by ∼5 h (Fig. 5A). As a control, lytA was interrupted to ensure autolysis was LytA dependent in both the Spn1+ and Spn1− strains. No autolysis was observed in the absence of LytA. In agreement with in vitro competition experiments (Fig. 3B), the Spn1+ strain outgrew the Spn1− strain; however, this effect was reversed in the absence of LytA (Fig. 5A). Genetic mutations in Mml had no effect on growth or autolysis (data not shown). We checked levels of LytA expression by Western blotting and detected no difference in expression of LytA between Spn1+ and Spn1− strains (Fig. 5B).

FIG 5.

FIG 5

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Spn1 has an effect on LytA-mediated autolysis. (A) Growth and autolysis of strain P1121 (Spn1+) and designated mutants in C+Y broth medium over 24 h at 37°C. Data are shown as means ± SEM. (B) Protein levels of LytA as determined by Western blotting on whole-cell lysates of Spn1+, Spn1−, and Spn1+ ΔlytA strains with anti-LytA antibody. Pneumolysin was used as a loading control. (C) Growth and autolysis of strain P1121 and designated mutants with and without LytA or Spn1 complemented by coculture in BHI for 16 h at 37°C. For each graph shown, red and green lines are strains grown in isolation. The blue line is the coculture of those two strains. Data are shown as means ± SEM.

To determine if Spn1 affected LytA activity, we complemented LytA activity by comparing growth of LytA+ and LytA− bacteria singly and in a coculture. If active, the LytA secreted from the LytA+ strain should be able to lyse the LytA− strain in coculture, resulting in a growth curve similar to that of an LytA+ strain grown in isolation. We measured the growth of a coculture of LytA+ and LytA− strains in both Spn1+ and Spn1− backgrounds. As expected, the growth pattern of the cocultured strains mimicked that of an LytA+ strain grown in isolation, indicating LytA is active and able to lyse LytA− strains in both the Spn1+ and the Spn1− backgrounds (Fig. 5C). This was confirmed by plating bacteria and colony counts at 0, 5, and 8 h (data not shown). To test if LytA's target, the cell wall, is altered in the Spn1+ strain, we cocultured the Spn1+ and Spn1− strains where both strains contained LytA. The growth curve mimicked the curve of Spn1+ strains grown in isolation (Fig. 5C). This led us to conclude that Spn1 confers resistance to LytA-mediated autolysis.

LytA has previously been associated with virulence (25). To determine if the delay in autolysis was the cause of the fitness defect in vivo, we competed Spn1+ ΔlytA and Spn1− ΔlytA strains in a 1:1 ratio in the murine model of colonization. The competitive advantage of the Spn1− strain was lytA independent (Fig. 3C). Thus, the altered cell wall physiology conferred by Spn1 may result in a fitness defect in vivo independent of its effects on promoting resistance to LytA-mediated autolysis.

Spn1 affects chain length.

LytA is also involved in cleavage of the cell wall to allow for cell separation during division. Deletion of lytA leads to the formation of long chains in some strains of pneumococci (26). Additionally, chain length has been shown to have an important role during both colonization and invasive disease (27, 28). Using bright-field microscopy, we observed that the mean chain length of the Spn1− strain was greater than that of the Spn1+ strain (Fig. 6A and B). The role of the phage-encoded homologue Mml in septation and chain length has not been elucidated. Surprisingly, deletion of lytA or mml did not lead to long chains in Spn1+ bacteria. However, Spn1+ ΔlytA Δmml bacteria did form long chains, a result consistent with the functional redundancy of these amidases. An alignment of the amino acid sequences of Mml and LytA showed that all catalytically active residues and those involved in dimerization are conserved, which is consistent with their functional redundancy (data not shown) (29, 30). Even when the effects of Mml are accounted for by comparing the Spn1− strain to the Spn1+ Δmml strain, the Spn1− strain grows as longer chains than the Spn1+ strain. The observations on chaining provided further evidence that the presence of Spn1 affects cell wall physiology.

FIG 6.

FIG 6

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Spn1 has an effect on the cell wall. (A) Images of strain P1121 and the mutant indicated in mid-log-phase culture using a bright-field microscope. The bar indicates 10 μm. (B) Average pixels per particle as determined by ImageJ analysis to quantify chain length. Student t test was used to compare groups. ***, P < 0.001. ns, nonsignificant. Data are shown as means ± SEM. (C) Growth of Spn1+ and Spn1− strains in 30 ng/ml of penicillin in tryptic soy broth for 5 h at 37°C. Data are shown as means ± SEM. (D) Competitive indexes (CI; output ratio/input ratio) of Spn1−/Spn1+ following growth for 5 h with increasing concentrations of penicillin. Each symbol is from a single culture. A Student t test was used to determine significant differences between groups. ***, P < 0.001.

To determine if the difference in chain length was causing the fitness defect in vivo, we assessed competitive colonization of the Spn1− ΔlytA strain and the Spn1+ ΔlytA Δmml strain, which both form long chains. Despite controlling for chain length, the in vivo fitness defect of the Spn1+ background remained (Fig. 3C).

Spn1 affects penicillin-mediated lysis.

Penicillin acts by inhibiting penicillin-binding proteins, which are required for cross-linking of the peptidoglycan backbone to prevent osmotic lysis (31). Strain P1121 is sensitive to penicillin (MIC, 50 ng/ml). However, to further demonstrate that Spn1 altered the pneumococcal cell wall, we examined whether there were small differences in penicillin sensitivity conferred by the Spn1 prophage. The MIC of the Spn1− strain was 10 ng/ml, indicating the presence of Spn1 resulted in a 5-fold higher MIC to penicillin. This effect was also demonstrated by differences in growth of Spn1+ and Spn1− strains at concentrations of penicillin near the MIC, 30 ng/ml (Fig. 6C). Additionally, we grew Spn1+ and Spn1− strains in coculture to determine differences in competitive growth in the presence of penicillin. At 10 ng/ml of penicillin, the competitive growth advantage of the Spn1+ strain is significantly increased (Fig. 6D). This difference in growth in the presence of penicillin was independent of LytA (data not shown).

DISCUSSION

Our findings demonstrate that expression of the phage-like genetic element Spn1 affects cell wall physiology and nasopharyngeal colonization of S. pneumoniae. Several aspects of this study merit further comment. Excision and integration of the Spn1 element occurs at the same site as that in the related phage MM1 (21). Through PCR analysis, it appears that a circularized form of the prophage occurs during normal bacterial growth, although we were unable to detect packaged phage by transmission electron microscopy or DNase protection assays. We were also unable to demonstrate phage infectivity by plaque assays (even after induction by mitomycin C), transduction, or detection of lysogenic conversion using a marked phage after coculture, in vitro or in vivo. Furthermore, we were unable to transfer phage Spn1 to other pneumococcal backgrounds, including TIGR4 and R36A. It remains possible that under different growth conditions or in its natural environment, Spn1 forms infectious phage particles. Spn1 could have become defective after infecting this strain, or it could have been obtained through transformation rather than infection. It was unexpected that the lytic genes of Spn1 were expressed during normal growth, yet we did not detect increased bacterial lysis compared to that of an isogenic strain lacking Spn1. The circularized form may be a precursor to Spn1 gene expression (and allow for amplification of gene expression), since mutations at both ends of the element in Spn1+ Δint Δmml prevented both circularization and expression of lytic and lysogenic genes. The inability of Spn1 to silence expression of lytic genes, which is not typical of lambdoid phages, appears to have required adaptation by the host bacterium (32).

The tolerance of prophage gene expression, even when further induced with mitomycin C, suggests that the phage lysins, which usually target the cell wall, are ineffective against strain P1121. It is possible that the lysins are inactive or only a small portion of the population is being activated by mitomycin C. However, our observations suggest that strain P1121 modifies its cell wall so that it is less susceptible to lysis. Several lines of evidence point to an alteration in cell wall physiology when this strain carries Spn1. The Spn1+ strain has a delayed autolysis phenotype not due to any alteration in amounts or activity of the major amidase responsible for autolysis, LytA. This suggests that LytA's target, the cell wall, is more resistant to its enzymatic activity. Moreover, the Spn1+ strain showed an increased resistance to penicillin. However, we did not detect an increased amount of peptidoglycan when purified from the Spn1+ strain or a thicker cell wall in electron micrographs of the Spn1+ strain (data not shown). The effect of penicillin suggests that alteration of the cell wall is due to increased cross-linking, since this is the step inhibited by the drug. Increased integrity through more extensive cross-linking within the cell wall could also explain the increased resistance to LytA and delayed autolysis, phenotypes previously linked to altered penicillin resistance (33, 34). Finally, alterations in cell wall physiology may be seen as differences in cell division/separation resulting in abnormal chain formation. The presence of Spn1 correlated with decreased average chain length.

Studies have shown that lytA mutants do not autolyze and in some strains display increased chain length (26). However, in this study, the Spn1+ strain demonstrated delayed autolysis, indicating resistance to LytA, but had shorter chains, which is usually linked to an increase in susceptibility to LytA. This apparent discrepancy could be indicative of differences in secretion and localization of amidases on the bacterial cell surface. A recent study on Mml and LytA showed that phosphorylcholine is required for secretion of both of these lysins. The authors of that study proposed that LytA binds the phosphorylcholine as it is added to the cell wall during division (35). This is supported by another study demonstrating that LytA is localized to the nascent peptidoglycan (33). This suggests that the change in the cell wall that leads to resistance to LytA in the Spn1+ strain occurs during maturation of the peptidoglycan, leaving the nascent peptidoglycan formed during division susceptible to LytA-mediated cleavage, resulting in reduced chain length. In contrast, despite conservation of catalytic amino acid residues between Mml and LytA, Mml was functionally redundant only in the chain length phenotype. This also may be due to localization of Mml at the nascent peptidoglycan and indicative of separate regulatory machinery for LytA secretion during autolysis.

The precise nature of the alteration in the pneumococcal cell wall required for tolerance of Spn1 is the subject of further investigation. Factors regulating cell wall synthesis, quantity of peptidoglycan per cell, and level of cross-linking are incompletely understood. These differences could be the direct action of an unknown protein on the prophage. However, it is possible that the overproduction of prophage products, including lysin and holins, leads to a general stress response in the host bacterium. Stress responses in bacteria often induce changes in the cell wall. For example, in Staphylococcus aureus there is evidence of cell wall thickening in the presence of vancomycin (36). Also, the CiaRH and WalRK two-component systems in S. pneumoniae are thought to maintain cell wall integrity in the presence of various stressors (37, 38).

The presence of Spn1 was associated with reduced fitness during competitive colonization, an effect that may be related to the changes in its cell wall. There are several phenotypes associated with the presence of Spn1 that may affect fitness during colonization. Most notably, greater chain length increases adherence to epithelial cells (28), and the presence and action of LytA is important during pathogenesis, although the exact mechanism is unclear (25). However, when we eliminate differences in chain length and the effects of LytA but maintain the presence of Spn1, we still observe a fitness defect in the Spn1+ strain during competitive colonization. This implies that the inherent differences in the cell wall, not these related phenotypes, are responsible for the fitness defect in vivo. Theoretically, alteration of the cell wall in response to the prophage could be a metabolic burden on the Spn1+ strain, which gives the Spn1− strain the edge in competitive colonization. The in vitro competition data support this concept, since the competitive effect is lost in a nutrient-rich environment. Additionally, the cell wall anchors many factors linked to fitness for colonization, which also could be playing a role (39).

The competitive fitness defect and alteration of the cell wall are both dependent on prophage gene expression. This indicates that a trans-acting factor encoded on the prophage is responsible for these phenotypes. We eliminated the potential contribution of three candidate trans-acting factors, Mml, PblB, and integrase. Mml has previously been shown to be redundant with LytA in phage progeny release, indicating it contributes to bacterial lysis (22). Integrase is a DNA binding protein involved in the excision and integration of the prophage (24). PblB was originally annotated on phages of Streptococcus mitis and is thought to mediate adherence to platelets (40). We saw no differences in adherence of the Spn1+ and Spn1− bacteria to A549 human epithelial cells in culture (data not shown). We also investigated the role of the putative holins encoded upstream of Mml. We found that deletion of the holins did not alter the phenotype of the Spn1+ strain in any in vitro assays and did not improve the fitness of the Spn1+ strain in vivo (data not shown). While we eliminated several potential trans-acting factors, there are ∼63 open reading frames annotated on Spn1, many of which have an unknown function and could be affecting bacterial fitness. The increased protein synthesis burden of carrying Spn1 also could account for the effect on fitness in vivo.

The question remains as to why the bacterium does not spontaneously cure Spn1 if it is associated with a fitness cost. S. pneumoniae is highly proficient at recombination and it should readily eliminate any disadvantageous genetic elements, yet Spn1 remains stably integrated (41). Moreover, in a study of human experimental pneumococcal colonization, Spn1 was present in the initial clinical isolate used to inoculate several volunteers and also in all of the isolates retrieved from participants up to 122 days later (10 and data not shown). The success of this lineage may be due to a factor found in the human host, such as exposure to penicillin, a natural population of phage, or competition with other pneumococci. The human host from which P1121 or its predecessor were isolated may have been exposed to penicillin, thereby selecting for the presence of Spn1. Also, colonization by multiple strains of pneumococci commonly occurs in children (42). Studies done in E. faecalis, a close relative of S. pneumoniae, show that lysogens are better able to compete against strains of the same species by way of phage lysis (43). Furthermore, prophages are known to protect against superinfection by other related phages. Therefore, it is possible that selective pressure from penicillin, other pneumococci, or other phages during colonization of a human host was sufficient to maintain this element.

In conclusion, we have determined that a novel prophage, Spn1, is associated with altering the bacterial cell wall and a fitness defect during competitive colonization. This type of interaction between a prophage and its bacterial host, where the bacteria tolerate the phage despite its negative effects, has, to our knowledge, not been described previously.

ACKNOWLEDGMENTS

This work was supported by grants from The National Institutes of Health to J.N.W. (R01 AI078538 and T32 A1060516) and the Morphology Core of the Center for Molecular Studies in Digestive and Liver Diseases (NIH P30 DK050306).

We also thank the Electron Microscopy Resource Laboratory.

Footnotes

Published ahead of print 9 May 2014

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