Strains used in this study are listed in **Table 1**. For routine growth, all bacterial strains were grown in Lysogeny broth (LB; 10 g of tryptone (Neogen, Lansing, Michigan, USA), 5 g of yeast extract (Neogen, Lansing, MI, USA) and 5 g of NaCl per liter) and incubated at 37◦C at 150 rpm for 5 h. The LB medium was solidified by addition of 1.5% agar (Neogen, Lansing, MI, USA) (Pasvolsky et al., 2014). Although, LB is suitable for bundle formation experiments, it was found to be less favorable for colony type biofilm or pellicle formation (Branda et al., 2001; Shemesh and Chai, 2013). Therefore, we studied colony biofilm and pellicle formation using chemically defined medium (CDM). CDM was prepared as previously described with slight modifications (Branda et al., 2001). Briefly, CDM contained 5mM potassium phosphate (pH 7), 100 mM 3-[*N*-Morpholino] propane sulfonic acid (MOPS) (pH 7), 2 mM MgCl2, 700 μM CaCl2, 50 μM MnCl2, 50 μM FeSO4, 1 μM ZnCl2, 2 μM thiamine (Sigma–Aldrich, St. Louise, MI, USA), 0.5% glycerol, 0.5% glutamate, 50 μg/ml tryptophan (Sigma–Aldrich, St. Louise, MI, USA), and 50 μg/ml phenylalanine. (Sigma–Aldrich, St. Louise, MI, USA). For CDA, 1.5% agar (Neogen, Lansing, MI, USA) was added. Medium and plates were freshly prepared and used the following day.

LBGM media was prepared as described previously by supplementing LB with 1% (v/v) glycerol and 0.1 mM MnSO4 (Shemesh and Chai, 2013).

### Lactose Preparation

A stock 50% lactose (w/v) (J. T. Baker, London, UK) solution was prepared in distilled deionized water and sterilized using a 0.2 μm filter (Whatman, Dassel, Germany). The stock solution of lactose was diluted in LB to final concentrations of 0–5% (w/v) or CDA to final concentration of 3% (w/v) (Assaf et al., 2015).

#### Biofilm Formation

Colony biofilms are produced when cells are placed on a solid agar surface. Importantly, one of the major characteristics of biofilm colony is the production of extracellular matrix which harbors the biofilm bacteria (Vlamakis et al., 2013). For colony type biofilm formation (Branda et al., 2001), starter cultures were prepared as describe above. 2.5 μl (around 3 × 105 CFU) from the starter culture was dropped on CDA with or without 3% lactose. The plates were incubated at 30◦C for 24 h. Images were taken using a Zeiss Stemi 2000-C microscope with an axiocamERc 5s camera.

For bundle formation, an overnight culture of cells was diluted 1:100 (to obtain O.D.(600) of 0.07) into LB supplemented with or without 3% lactose (w/v). The samples were incubated at 37◦C at 50 rpm for 5 h (O.D.(600) of 1). One milliliter of each sample was collected and centrifuged at 5000 rpm for 2 min. The supernatant was discarded, the pellet was resuspended and 3 μl of the suspension placed on a glass slide was visualized in a transmitted light microscope using Nomarski differential interference contrast (DIC), at 40× magnification

#### TABLE 1 | Strains used in this study.

(Pasvolsky et al., 2014; Oknin et al., 2015). Furthermore, a confocal laser scanning microscope (CLSM) was used to visualize cyan fluorescent protein (CFP) or green fluorescent protein (GFP) expression. CFP expression of strain YC189 was observed using 458-nm argon laser, while GFP expression of strains YC161 and YC164 was observed using 488-nm argon laser (Zeiss LSM510 CLS microscope, Carl Zeiss, Oberkochen, Germany).

For pellicle formation, bacteria were inoculated from the agar plates into LB broth and incubated for 5 h at 37◦C at 150 rpm. Next, 5 μl of the culture was seeded in a 12 wells plate (Nunc, Roskild, Denmark) containing 4 ml of CDM per well. The plates were incubated at 30◦C. Pictures were taken after 24 h using SAMSUNG Galaxy camera.

#### AI-2 Production Assay

To determine the effect of lactose on AI-2 production, we used a bioluminescence assay as described before (Aharoni et al., 2008; Shemesh et al., 2010). Briefly, *B. subtilis* cells were grown in conditions inducing bundle formation as described above. One milliliter of each sample was collected and centrifuged at 5000 rpm for 2 min. Supernatant was collected and neutralized to pH 7 using 1 M NaOH. An overnight culture of *Vibrio harveyi* MM77, a mutant strain which does not produce either AI-1 nor AI-2, was diluted 1:5,000 in a mixture of 90% (v/v) fresh AB medium and 10% (v/v) neutralized supernatant to a total volume of 200 μl per well. The negative control contained bacteria in fresh AB medium alone, while the positive control contained the bacteria, fresh AB medium and 10% (v/v) supernatant medium containing AI-2 of *V. harveyi* BB152 (AI-1–, AI-2+). The luminescence readings were performed in triplicate in white 96-well plates with an optic bottom (Nunc, Roskild, Denmark) using a plate reader (GENiosTECAN, NEOTEC Scientific Instrumentation Ltd. Camspec, Cambridge, UK) at 30◦C. Luminescence measurements were recorded every 30 min in parallel with optical density (595 nm) readings. To avoid dissimilarities caused by differences in growth rates, the relative luminescence (RLU) was calculated. Briefly, the value of each reading was divided by the optical density values to normalize the luminescence value of each sample to its cell density. Fold induction above the nonspecific luminescence background of the negative control was determined at the end of bacterial growth, after approximately 20 h of growth. The area under the curve (AUC) was calculated to better demonstrate the differential expression in RLU values by means of the sum of: the average of *Y* values/the average of *X* values (Aharoni et al., 2008; Soni et al., 2015).

### AI-2 Effect on Biofilm Formation

To determine the effect of AI-2 on bundle formation as well as *tapA* expression, we used (S)-4,5-Dihydroxy-2,3 pentandione (DPD) (Omm Scientific, Inc, Dallas, TX, USA) which is the precursor for AI-2. Bacterial cells prepared as described above and were incubated in the presence of DPD in LB at 37◦C at 50 rpm for 5 h. The cells were collected and visualized in a transmitted light microscope using DIC. Furthermore, a CLSM was used to visualize CFP expression using 458-nm argon laser (Oknin et al., 2015). For complementation tests, DPD was supplemented in LB medium to final concentration of 200 μM as an exogenous precursor for AI-2.

#### Statistical Analysis

The data obtained were analyzed statistically by means of ANOVA following *post hoc t*-test with Bonferroni correction using Microsoft Excel software. *P*-values less than 0.01 were considered significant.

### RESULTS

#### Lactose Induces Biofilm Formation by *B. subtilis*

Initially, we found that addition of lactose to growth media such as LB or chemical defined agar (CDA) enhances biofilm

formation by *B. subtilis*. As it can be seen in **Figure 1A**, a majority of *B. subtilis* (YC161) cells preferably generated long chains of cells attaching to each other to form a biofilm-related structure (bundle) in the presence of lactose. Similarly, lactose also induced colony type biofilm formation on CDA, as seen in the center of the colony (**Figure 1B**). The structure of the biofilm formed on the CDA with addition of lactose has higher structure complexity. Accordingly, the morphology of the biofilm in the presence of lactose is more developed and structured as seen in the center of the colony (**Figure 1B**). Subsequently, we tested whether the increase in biofilm formation in the presence of lactose is due to the increase in bacterial growth rate. The bacterial growth of *B. subtilis* was not affected by addition of lactose (Supplementary Figure S1). Therefore, we assume that the effect of lactose is specific to the biofilm formation.

## Lactose Up-Regulates Expression of Genes Associated with Extracellular Matrix Production

In order to confirm our above findings and to determine if the bundles induced by lactose are biofilm related, we used genetically modified *B. subtilis* strains, which express fluorescent proteins under the control of important extracellular matrix related promoters. To examine the expression of *tapA* operon, we used the strain (YC189) which produces CFP under the control of the *tapA* promoter, whereas, the expression of *eps* operon was determined using strain (YC164) which produces GFP under the control of *eps* promoter (Chai et al., 2008). The amounts of the fluorescent proteins as well as their intensity represent the expression of the tested promoter in the different samples. As it is demonstrated in **Figure 2**, the expression of both operons was increased as a result of lactose introduction into the growth medium. Moreover, mutant strains which are defected in production of extracellular matrix showed deficiency in bundles formation in the presence of lactose (**Figure 3**).

## Lactose Triggers AI-2 Production

Next, we decided to test whether lactose affects AI-2 production. Using *V. harveyi* MM77 as a reporter strain enables us to examine the effect of lactose on QS via the LuxS dependent pathway. Supernatants from *B. subtilis*, grown with or without lactose, were used for evaluating the amount of AI-2 secreted to the media. The RLU indicates the relative amount of AI-2 in the suspension; a peak of the relative bioluminescence was detected following 14 h in all tested samples which was found to be remarkably higher in the presence of lactose (in dose dependent manners; **Figure 4A**). Indeed, there was a significantly increase in the production of AI-2 by *B. subtilis* cells in the presence of all tested lactose concentrations especially in the presence of 3% of lactose (**Figure 4B**).

## *luxS* is Essential for Biofilm Formation in the Presence of Lactose

We further investigated the connection between LuxS dependent QS and induction in biofilm formation. Thus, we used the YC189 strain (harboring the P*tapA-cfp* transcriptional fusion) which was grown in the presence of different concentrations of DPD (precursor for AI-2). Interestingly, increasing concentrations of DPD stimulated the biofilm bundles formation as well as *tapA* expression (**Figure 5**). The induction in bundle formation and *tapA* expression seems to be in linear correlation with the concentration of DPD.

To further investigate a possible role of LuxS on biofilm formation in the presence of lactose, we tested the ability of *B. subtilis luxS* mutant to form bundle as well as pellicle and colony biofilm with or without lactose. As seen in **Figure 6**, the *luxS* mutant is somehow defected in generating developed and structured pellicle and colony biofilm in the presence of lactose compared to the WT. Furthermore, *luxS* mutant could not form biofilm bundles in the presence of lactose (**Figure 7**). Interestingly, addition of DPD restored at least partially the bundling phenotype of the *luxS* mutant (**Figure 7**).

#### DISCUSSION

Our results show that lactose triggers bundle formation as well as formation of colony type biofilm by *B. subtilis.* This result falls in line with our previous study which showed that lactose enhances biofilm formation by *Streptococcus mutans* (Assaf et al., 2015). Expression of *epsA-O* and *tapA* operons, which are responsible for biofilm matrix

production, were notably increased when lactose was added to the LB medium (**Figure 2**). Interestingly, induction in expression of both operons is correlated with biofilm bundles formation by *B. subtilis* cells. Bundle formation is one of the first stages in biofilm development (Branda et al., 2001). Moreover, investigation of the mutant strains for these operons shows absence of the bundling phenotype as a response to lactose (**Figure 3**). This result indicates that lactose induce biofilm formation depends on *tapA* and *epsA-O* expression.

In recent years, there has been an increasing interest in the quorum-sensing signaling molecules related to food spoilage. Various signaling compounds associated with QS, such as AI-2, have been detected in different food systems such as milk (Pinto et al., 2007). Furthermore, studies have shown that QS is important for social behavior of *B. subtilis* and other bacteria (Lombardía et al., 2006). Using *V. harveyi* as a reporter strain for bioluminescence, we were able to track the level of produced AI-2 molecules. We observed an increase in the AI-2 production as a response to lactose in dose dependent manners (**Figure 4**). It has been shown previously that simple dietary sugars can affect QS, specifically production of AI-2 by *Klebsiella pneumoniae* (Zhu et al., 2012). In our study, the cell density of all tested samples was the same at the sampling time, consequently, changes in the AI-2 production is apparently not related to the cell density but to the metabolic state of the bacteria. Thus, our results support previous studies that showed that AI-2-dependent signaling is a reflection of metabolic state of the cell and environmental factors and not cell density (Bassler, 1999; Beeston and Surette, 2002). Previous studies also suggested that activation of QS through LuxS can be regulated in response to sugar metabolism by cyclic AMP receptor protein molecules (Lyell et al., 2013). In *B. subtilis* cells, lactose may affect the energetic metabolic balance in the cell, and through second messengers such as cyclic AMP, or CCP can lead to expression of QS genes such as *luxS*.

The main finding of this study is the apparent link between lactose induced biofilm formation and activation of QS system through increased production of AI-2 molecules in *B. subtilis*. Addition of synthetic precursor for AI-2, DPD, to the media resulted in enhanced bundle formation as well as up-regulation

were taken using a Zeiss Stemi 2000-C microscope with an axiocamERc 5s camera. Images are representative of four biological repeats. (B) WT and *luxS* cells were used for pellicle biofilm formation. Biofilms were generated in chemical defined medium (CDM) and CDM supplemented with 3% lactose. Pictures were taken using Sumsung galaxy digital camera. Images are representative of two biological repeats.

of *tapA* expression (**Figure 5**). Similarly, the direct effect of AI-2 molecules on EPS biosynthesis has been observed previously in *Vibrio cholera*e where the AI-2 molecules up-regulated expression of the EPS biosynthesis genes (Hammer and Bassler, 2003). According to our results, examination of biofilm formation in CDM of the *B. subtilis luxS* mutant resulted in deficiency of biofilm formation (bundle, and colony types) (**Figures 6A** and **7**). These results suggested that QS via LuxS cascade plays an important role in biofilm formation in the presence of lactose. This is consistent with previous research which showed that LuxS is important for *B. subtilis* social behavior (motility and biofilm formation) (Lombardía et al., 2006). Another study showed that blocking the AI-2 pathway, using an AI-2 analog, inhibited biofilm formation by *B. subtilis* (Ren et al., 2002). Similar results were found for *Hafnia alvei*, a foodrelated bacterium that can be found in dairy products. QS in *H. alvei* is required for differentiation of individual cells into a complex multicellular structure of biofilm (Souza Viana et al., 2009).

Interestingly, we observed that the *luxS* mutant strain could form pellicle in biofilm promoting medium LBGM (**Figure 8**). Although, a pellicle formation in LBGM appears to be not LuxS dependent, it seems that in CDM there is a slight induction in pellicle formation in response to lactose (**Figure 6B**). As it was shown recently (Shemesh and Chai, 2013), glycerol and manganese activate KinD-Spo0A pathway for matrix production. In case of lactose, it seems that enhanced production of AI-2 affects not directly on the biofilm formation cascade. This may explain the differences found between phenotypes in CDM supplemented with lactose and in LBGM. Activation of biofilm formation via QS system might be an additional regulatory mechanism which enables fine tuning of the biofilm formation pathway that has been previously described (Shemesh and Chai, 2013).

The LuxS system possesses an inherent metabolic function in the activated methyl cycle; phenotypic defects in *luxS* mutants may not strictly be attributed to AI-2 signaling but possibly to metabolic disturbances. For instance, biofilm defects in a *Lactobacillus rhamnosus luxS* mutant are not restored by AI-2 molecules but rather by the addition of cysteine, indicating a sole metabolic role of LuxS (Lebeer et al., 2007). In order to test whether the deficiency of biofilm formation in the presence of lactose in the mutant strain is due to AI-2 signal molecules or due to metabolic reason, we used DPD for complementation tests. It was shown previously that the synthetic AI-2 precursor (DPD) has the ability for specific AI-2 complementation during biofilm formation by *Streptococcus intermedius* (Ahmed et al., 2008). In the complementation test, we observed restoration of the biofilm phenotype. The *luxS* mutant showed ability for increased bundle formation in media supplemented with lactose and 200 μM of DPD (**Figure 7**), indicating that the abolished biofilm formation is mostly connected to AI-2 and not to LuxS enzyme function.

In overall, results of the present study suggest that QS via LuxS system plays an important role in biofilm formation induced by lactose in *B. subtilis*. As lactose affects activation of LuxS system, it is likely related to activation of Spo0A which leads to biofilm formation through a known pathway of up-regulation of the extracellular matrix operons. Moreover, Spo0A has been shown to be a negative regulator of LuxS system (Lombardía et al., 2006). Additional research on lactose in association with QS will further elucidate the role of QS in biofilm formation of *Bacilli* and the effect of this dairy component on biofilm related gene expression.

#### REFERENCES

#### AUTHOR CONTRIBUTIONS

DD-A together with MS planned the experiments and wrote the original manuscript. DD-A performed the experiments described in the manuscript. DS and YC assisted in planning biofilm experiments as well as revised the manuscript critically for important intellectual content. DD-A, DS, and MS integrated all of the data throughout the study and crafted the final manuscript.

#### ACKNOWLEDGMENTS

Contribution from the Agricultural Research Organization (ARO), the Volcani Center, Beit Dagan, Israel, No. 733/15-E Series is acknowledged. This work was partially supported by the COST ACTION FA1202 BacFoodNet and by the Israel Dairy Board grant 421-0270-15. DD-A is recipient of Scholarship of Excellency for outstanding Ph.D. students from The ARO.

#### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fmicb. 2015.01517

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Copyright © 2016 Duanis-Assaf, Steinberg, Chai and Shemesh. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# *Campylobacter jejuni* biofilms contain extracellular DNA and are sensitive to DNase I treatment

Helen L. Brown1, 2, Kate Hanman1 , Mark Reuter 1 , Roy P. Betts 3 and Arnoud H. M. van Vliet 1 \*

*1 Gut Health and Food Safety Programme, Institute of Food Research, Norwich, UK, 2 Cardiff School of Health Sciences, Cardiff Metropolitan University, Cardiff, UK, 3 Campden BRI, Chipping Campden, UK*

#### *Edited by:*

*Romain Briandet, Institut National de la Recherche Agronomique, France*

#### *Reviewed by:*

*Odile Tresse, French National Institute for Agricultural Research/Nantes-Atlantic National College of Veterinary Medicine, Food Science and Engineering, France Rikke Louise Meyer, Aarhus University, Denmark*

#### *\*Correspondence:*

*Arnoud H. M. van Vliet, Institute of Food Research, Norwich Research Park, Colney Lane, Norwich NR4 7UA, UK<EMAIL_ADDRESS>
#### *Specialty section:*

*This article was submitted to Food Microbiology, a section of the journal Frontiers in Microbiology*

*Received: 04 March 2015 Accepted: 26 June 2015 Published: 10 July 2015*

#### *Citation:*

*Brown HL, Hanman K, Reuter M, Betts RP and van Vliet AHM (2015) Campylobacter jejuni biofilms contain extracellular DNA and are sensitive to DNase I treatment. Front. Microbiol. 6:699. doi: 10.3389/fmicb.2015.00699*

Frontiers in Microbiology | www.frontiersin.org July 2015 | Volume 6 | Article 699 |

Biofilms make an important contribution to survival and transmission of bacterial pathogens in the food chain. The human pathogen *Campylobacter jejuni* is known to form biofilms *in vitro* in food chain-relevant conditions, but the exact roles and composition of the extracellular matrix are still not clear. Extracellular DNA has been found in many bacterial biofilms and can be a major component of the extracellular matrix. Here we show that extracellular DNA is also an important component of the *C. jejuni* biofilm when attached to stainless steel surfaces, in aerobic conditions and on conditioned surfaces. Degradation of extracellular DNA by exogenous addition of DNase I led to rapid biofilm removal, without loss of *C. jejuni* viability. Following treatment of a surface with DNase I, *C. jejuni* was unable to re-establish a biofilm population within 48 h. Similar results were obtained by digesting extracellular DNA with restriction enzymes, suggesting the need for high molecular weight DNA. Addition of *C. jejuni* genomic DNA containing an antibiotic resistance marker resulted in transfer of the antibiotic resistance marker to susceptible cells in the biofilm, presumably by natural transformation. Taken together, this suggest that eDNA is not only an important component of *C. jejuni* biofilms and subsequent food chain survival of *C. jejuni*, but may also contribute to the spread of antimicrobial resistance in *C. jejuni*. The degradation of extracellular DNA with enzymes such as DNase I is a rapid method to remove *C. jejuni* biofilms, and is likely to potentiate the activity of antimicrobial treatments and thus synergistically aid disinfection treatments.

Keywords: *Campylobacter jejuni*, biofilm, food safety, extracellular matrix, extracellular DNA, antibiotic resistance

#### Introduction

Campylobacter jejuni is the most common cause of bacterial foodborne infection within the UK (Nichols et al., 2012). Its success as foodborne pathogen contrasts with its fastidious nature, as it requires specific atmospheric conditions, nutrient-rich growth medium and has a narrow temperature range (between 35 and 45◦C) for growth. Several mechanisms for survival in the food chain have been proposed, including the ability of C. jejuni to enter a viable but none culturable (VBNC) state (Rollins and Colwell, 1986), as well as formation of de novo biofilms or integration into existing (multispecies) biofilms (Teh et al., 2014). Biofilms are defined as surface attached populations, either single or multiple species, which are surrounded by a self-produced extracellular matrix (Donlan, 2002). The extracellular matrix differs depending on the species within the biofilm but typically comprises of DNA, proteins and polysaccharides (Branda et al., 2005).

The extracellular matrix is an essential component of bacterial biofilms, and usually accounts for more than 90% of the dry mass of a biofilm (Flemming and Wingender, 2010). It allows cells to remain hydrated and metabolically active by trapping nutrients and liquid near the bacterial cells. It also reduces access of larger molecules such as antimicrobials (Mulcahy et al., 2008; Billings et al., 2013), leading to increased bacterial persistence, and is structurally important, maintaining the shape of the biofilm and ensuring the cohesion of the biofilm (Sutherland, 2001). Extracellular DNA (eDNA) appears to have a structural role in the biofilms of many different species, including Pseudomonas aeruginosa (Chiang et al., 2013), Staphylococcus aureus (Mann et al., 2009), Listeria monocytogenes (Harmsen et al., 2010), and Escherichia coli (Zhao et al., 2013).

Recent studies have shown that eDNA is important for biofilm establishment and maintenance by C. jejuni strain 81–176 in laboratory conditions (Bae et al., 2014; Svensson et al., 2014), but this has not yet been studied in the context of the conditions encountered by C. jejuni in the processing environment. Previous studies have shown that food chain relevant conditions such as atmospheric oxygen levels (Reuter et al., 2010), reduced temperatures (Buswell et al., 1998) and surface soiling (Brown et al., 2014) all increase C. jejuni biofilm formation, and as such may also influence the composition of the C. jejuni biofilm, necessitating the study of C. jejuni biofilms in these conditions.

The aim of this study was to further investigate the role of eDNA within the C. jejuni biofilm, with particular reference to its role in food chain relevant environments. Here we present evidence that eDNA is also present in biofilms of C. jejuni reference strains NCTC 11168 and 81116 when incubated in aerobic conditions and on food chain relevant surfaces such as stainless steel. Degradation of eDNA by DNase I leads to a rapid loss of biofilm structure, releasing cells into the planktonic phase. Treatment of surfaces with DNase I also inhibits de novo biofilm formation, either due to re-growth from single, attached, cells or from de novo attachment of C. jejuni cells. Addition of C. jejuni DNA to biofilms results in the transfer of genetic markers, which can contribute to spread of antimicrobial resistance in C. jejuni populations.

#### Materials and Methods

#### *C. jejuni* Strains and Growth Conditions

C. jejuni strains NCTC 11168 (Parkhill et al., 2000), its derivative expressing a green fluorescent protein and chloramphenicol resistance marker (C. jejuni NCTC 11168 cj0046::gfp-CmR ) (Brown et al., 2015), strain 81116 (Pearson et al., 2007) and all microaerobic biofilm incubations were routinely cultured in a MACS-MG-1000 controlled atmosphere cabinet (Don Whitley Scientific) under microaerobic conditions (85% N2, 5% O2, 10% CO2) at 37◦C. For growth on plates, strains were either grown on Brucella agar or BAB with Skirrow antibiotic supplement (10µg/ml vancomycin, 5µg/ml trimethoprim, 2.5 IU polymyxin-B). Broth culture was carried out in Brucella broth (Becton & Dickinson).

#### *Campylobacter* Growth for Biofilm Assay

Frozen 50µl single-use glycerol stocks were thawed, inoculated onto Skirrow plates and grown overnight at 37◦C in microaerobic conditions (5% O2, 10% CO2 and 85% N2). Cells from the Skirrow plate were used to inoculate Brucella broth then grown overnight as a shaking culture (37◦C, microaerobic conditions). Following overnight growth, cell cultures were adjusted to an A600 of 0.05 in Brucella medium or Brucella medium supplemented with 5% v/v chicken juice. To allow biofilm formation, 1 ml of this solution was added to either a sterile borosilicate glass test tube (Corning) or 3 ml to a six-well polystyrene tissue culture plate (Corning) containing a sterile stainless steel coupon (Stainless steel type 1.4301 according to EN 10088-1, with a Type 2B finish according to EN 10088-2). In each biofilm assay a test tube containing sterile Brucella medium was incubated alongside the C. jejuni containing tubes to ensure sterility was maintained and, following crystal violet staining, to quantification of staining levels where biofilm was not present. Tubes were incubated at 37◦C in atmospheric air conditions using an Innova 4230 (New Brunswick Scientific) incubator at 37◦C. Unless otherwise stated all biofilms were formed in aerobic conditions at 37◦C for 48 h before staining procedures were carried out. For each assay a microaerobic biofilm control was also undertaken, to ensure that oxygen availability does not have a major effect on results and to allow comparison with previous studies (Reuter et al., 2010; Brown et al., 2013, 2014, 2015) . This sample was prepared in exactly the same way as the aerobic biofilm cultures but test tubes were placed back in the 37◦C microaerobic incubator for all static incubations.

#### Preparation of Chicken Juice

Chicken juice was prepared as described previously (Brown et al., 2013, 2014). Briefly, frozen commercially available whole chickens were purchased from UK supermarkets before thawing at room temperature. Exudate was collected, centrifuged to remove debris and sterilized by using a 0.2µm sterile polyethersulfone (PES) syringe filter (Millipore) before aliquotting and storage at −20◦C until use. Chicken juice was diluted v/v in Brucella medium for use in biofilm assays.

#### Enzyme Treatment of *C. jejuni* Biofilms

For DNase I treatments, unless otherwise stated, a volume of 4µl DNase I enzyme (Fermentas), giving a final concentration within the biofilm of 4 U/ml v/v and 4µl of DNase I buffer (Fermentas) were added to each test tube, along with 1 ml of diluted cell suspension at either the start of the static incubation or after 12, 24, 36, or 48 h of static incubation. Following treatment, static cultures were placed back in 37◦C, aerobic conditions to complete the 48 h incubation before staining with crystal violet to allow biofilm quantification. For restriction enzyme digest of biofilms 4µl of 10 U/µl BamHI, BlpI, HaeIII, HindIII, MscI or RsaI, (NEB), or DNase I (Fermentas), or RNase (QIAGEN) were added to test tubes containing diluted C. jejuni suspension prior to static incubation and then incubated at 37◦C for 48 h in aerobic conditions. Equal volumes (4µl) of the buffers and bovine serum albumin were also added if recommended by the manufacturers. For the assays assessing the time required for DNase I activity, biofilms were allowed to form for 48 h before addition of 4 U/ml v/v DNase I enzyme (1 U/µl, Fermentas) and 4µl of DNase I buffer to the samples, followed by incubation for up to 2 h. During the incubation with enzyme the samples were placed in 37◦C, aerobic conditions. All samples were subsequently stained with crystal violet.

For assessment of biofilm regrowth, biofilms were allowed to establish for 48 h followed by a 15 min incubation with DNase I. Tubes were then washed twice with sterile PBS followed by addition of either an equal volume of bacterial culture with an A600 of 0.05, or sterile Brucella medium, followed by a further 48 h incubation at 37◦C in aerobic conditions. All samples were subsequently stained with crystal violet. In order to ensure consistency between control and treatment samples all tubes were manipulated in exactly the same way, being removed and placed back in the same incubation conditions during each enzyme addition. Heat inactivated DNase I was prepared by incubating an aliquot of DNase I and its buffer at 95◦C for 10 min and allowing to cool before addition to the biofilm cultures.

#### Visualization of Extracellular DNA from Shaking Cultures and Biofilms

Following incubation to allow biofilm formation in both aerobic and microaerobic conditions, the supernatant was removed and the tubes were rinsed once with sterile PBS to remove loosely attached bacterial populations. After rinsing and removal of the rinse suspension a second 1 ml volume of sterile PBS was added to each test tube and a sterile cotton wool swab was used to gently swab to walls of the test tube, releasing the biofilm from the walls of the test tube and in to suspension. The resuspended biofilm (PBS containing the loosened biofilm cells) and supernatant (liquid initially contained within the test tube) from several biofilm cultures were collected and pooled before diluting to a A600 of 0.3. Aliquots were mixed with gel loading buffer (NEB) and added to a 0.9% agarose gel and run at 100 V for 45 min in 0.5% TBE buffer. A 1 kb ladder (NEB) was used for size comparison. Following electrophoresis, nucleic acids were stained using ethidium bromide, and DNA was visualized using a GelVue UV light and documented using a U:Genius gel documentation system (Syngene). The amount of eDNA in planktonic and biofilm fractions was quantified by comparing the intensity of the DNA bands after UV illumination and comparison with the 3 kb marker fragment (125 ng), using ImageJ (Rasband, W.S., ImageJ, U. S. National Institutes of Health, Bethesda, Maryland, USA, http://imagej.nih.gov/ij/, 1997–2014). Quantification was based on three independent experiments.

#### Restriction Digest of *C. jejuni* Genomic DNA

A 1µl volume of restriction enzyme (BamHI, BlpI, HaeIII, HindIII, MscI, or RsaI, all supplied by NEB), or DNase I (1U/ µl, Fermentas), or RNase (QIAGEN) was added to a mixture containing ∼500 ng of C. jejuni NCTC 11168 or 81116 genomic DNA, prepared using a commercial kit (QIAGEN) following manufactures guidelines, 1µl of 10× enzyme buffer (if required), 1µl of 1 mg/ml BSA (if required) and molecular grade water to a final volume of 10µl. Samples were incubated for 60 min in a 37◦C water bath to allow digestion of the genomic DNA. DNA was visualized using a GelVue UV light and documented using a U:Genius gel documentation system (Syngene).

#### Assessment of Natural Transformation within the Biofilm

Genomic DNA was extracted from the C. jejuni NCTC 11168 cj0046::gfp−CmR mutant (Brown et al., 2015) using a commercial kit (QIAGEN), following manufacturers guidelines. DNA concentration was calculated after the final elution and stored at −20◦C until use. The standard 48 h static biofilm incubation was carried out, using duplicate test tubes for all conditions. A total of 2µg genomic DNA was added to test tubes either prior to the start of biofilm incubation, or following 24 h of static incubation. Following a total of 48 h of incubation one test tube of each condition was stained using crystal violet and the second tube washed once with 1 ml PBS and the biofilm population released by swabbing with a sterile cotton bud. Both the supernatant and released biofilm population were retained for viability assessment.

#### Crystal Violet Staining

Cell suspensions were removed from the test tubes before washing with distilled water before drying at 60◦C for 30 min. A 1 ml of 1% w/v crystal violet solution was added and tubes were further incubated on a rocker at room temperature for 30 min. After this incubation, the non-bound dye was removed from the tubes by thorough washing in water followed by drying at 37◦C. Bound crystal violet was dissolved by adding 20% acetone/80% ethanol and incubating on a rocking platform for 15 min at room temperature. The resulting dissolved dye was measured at a wavelength of 590 nm using a Biomate 5 spectrophotometer (Thermo Scientific).

#### 2,3,5 Triphenyltetrazolium Chloride (TTC) Staining

This method was carried out as previously described (Brown et al., 2013, 2014). Following a 48 h incubation to allow biofilm formation, cell suspensions were removed and test tubes were washed twice with 1 ml of sterile PBS. A 1.2 ml volume of Brucella broth supplemented with 0.05% w/v TTC was then added to each test tube before further incubation at 37◦C in microaerobic conditions for 72 h. Following secondary incubation, the TTC solution was removed and the test tubes were air dried. Bound TTC dye was dissolved as above using 20% acetone/80% ethanol and the A500 of the solution measured.

#### Assessment of Cell Viability by Culture

To determine the number of viable cells, the planktonic fraction, or released biofilm population was 10-fold serially diluted eight times in PBS and 5µl of each dilution spotted on Brucella agar plates or (for assessment of natural transformation) Brucella agar containing 10µl/ml chloramphenicol. After 2 days of growth at 37◦C in microaerobic conditions, the dilution resulting in two or more colonies was recorded. Cell viability in biofilm assays was assessed upon initial addition of cultures into static culture and following static incubation, prior to crystal violet staining and, where necessary, following the 72 h TTC incubation.

#### Statistics

Statistical analysis was carried out using GraphPad Prism software. At least three biological replicates (each with three technical replicates unless otherwise stated) were used to calculate mean and standard deviation. Significance was measured using either a Mann–Whitney test (biofilm formation) or ANOVA (DNA yield).

#### Results

#### Extracellular DNA Is Present within the *C. jejuni* Biofilm and during Both Aerobic and Microaerobic Incubation

Biofilms of C. jejuni NCTC 11168 and 81116 were generated and used for the investigation of eDNA. Separation of nucleic acids on agarose gels showed the presence of extracellular DNA in both the biofilm and planktonic fractions, independent of whether the biofilm samples were incubated in aerobic or microaerobic conditions (**Figure 1**). Within the biofilm samples, there was no distinguishable difference between the eDNA bands produced by C. jejuni NCTC 11168 and 81116, although in the supernatant, C. jejuni NCTC 11168 cultures contained less DNA than C. jejuni 81116. The atmospheric conditions used for the incubation did not seem to affect eDNA levels, although as previously reported, total biofilm mass increased during aerobic biofilm incubation (Reuter et al., 2010; Brown et al., 2014).

#### Addition of DNase I Leads to Rapid Reduction of Biofilm Levels and Prevents Formation of New Biofilms

To assess whether the role of eDNA differs between different stages of biofilm maturity in C. jejuni, DNase I was added at 12 h intervals over the total of a 48 h incubation in aerobic conditions. There was no detectable C. jejuni biofilm after incubation with 4 U/ml DNase I, regardless of the age of the biofilm (**Figure 2A**), indicating that eDNA is an important extracellular matrix component during both initial attachment and maturation. We next assessed how rapidly degradation occurs by treating biofilms grown for 48 h with DNase I followed by detection with crystal violet at timed intervals over a two h period. Following only a 5 min incubation with DNase I, there was no detectable staining on the glass surface, and A590 values were indistinguishable from the negative control containing Brucella medium only (**Figure 2B**). Levels of staining did not reduce further at later time points, suggesting that a 5 min treatment can achieve degradation of the eDNA in the C. jejuni biofilm and results in a reduction of biofilm levels below the detection limit (Tresse et al., 2006).

Finally, the concentration of DNase I required to degrade the biofilm was also investigated. Addition of DNase I at concentrations ranging from 0.01 to 5 U/ml were significantly reduced crystal violet staining, and there was no statistically significant difference between DNase I treated test tubes and the negative control tube containing Brucella medium only (**Figure 2C**). It is interesting to note that DNase I treatment had no impact on cell viability, and most likely only degrades the biofilm matrix, resulting in the release of attached cells into

suspension. Biofilms incubated with DNase I in microaerobic conditions also showed the same pattern, confirming that the effects observed were not a response to atmospheric condition, but DNase I treatment. Inactivation of DNase I by heat treatment removed its ability to affect C. jejuni biofilms (Figure S1), but did not inhibit growth of C. jejuni.

The long-term effects of DNase I-mediated degradation of C. jejuni biofilms from abiotic surfaces was assessed by adding fresh C. jejuni NCTC 11168 culture to DNase I-treated and washed borosilicate test tubes previously containing a C. jejuni biofilm. There was no detectable C. jejuni biofilm in either the tubes with added Brucella medium or the tubes with added

FIGURE 2 | DNase I is able to rapidly degrade *C. jejuni* NCTC 11168 biofilms. (A) DNase I (4 units/ml) was added at defined intervals to aerobically incubated NCTC 11168 cultures over a 48 h static incubation and biofilm degradation assessed by crystal violet staining. (B) Following a 48 h static incubation to allow biofilm formation, DNase I was added to biofilms for between 5 and 120 min before biofilm degradation was assessed. (C) The concentration of DNase I required for biofilm control was also assessed using DNase I concentrations of between 0.01 and 5 U/ml. In each graph, "11168" represents an untreated biofilm culture of *C. jejuni* NCTC 11168 and "control" represents a tube containing sterile Brucella medium only. Error bars show standard deviation. Statistically significant results, as determined using the Mann–Whitney U test, are indicated using an asterisk (\**P* < 0.05, \*\**P* < 0.01, \*\*\**P* < 0.001).

Brown et al. eDNA and *C. jejuni* biofilms

C. jejuni in either aerobic or microaerobic conditions (**Figure 3**). This suggests that DNase I treatment is not only a rapid method of degrading C. jejuni NCTC 1168 biofilms but also prevents biofilm regrowth.

#### Restriction Digestion of eDNA Leads to Reduced Levels of *C. jejuni* Biofilm

The eDNA found within the C. jejuni NCTC 11168 and 81116 biofilms is of high molecular weight (**Figure 1**), and we speculated that biofilm formation requires high molecular weight nucleic acids, rather than simply the presence of any nucleic acids. Six restriction enzymes were selected, which are predicted to digest C. jejuni genomic DNA to a range of fragment sizes (**Figures 4C,D**), and these enzymes were assessed for their ability to degrade 48 h old C. jejuni biofilms. With C. jejuni NCTC 11168 there was a significant reduction in crystal violet staining for all six restriction enzymes tested, with little variation between enzyme treatment and the negative control (**Figure 4A**). Although the same trend was observed with C. jejuni 81116 biofilms, this was not statistically significant except for DNase I treatment (**Figure 4B**). This was consistent with the reduced digestion observed with C. jejuni 81116 genomic DNA, producing fragments of higher molecular weight than those obtained by digestion of C. jejuni NCTC 11168 genomic DNA (**Figure 4D**).

#### DNase I Treatment is Also Effective on Food Chain Relevant Surfaces

The effect of DNase I treatment on C. jejuni biofilms formed on food-relevant surfaces such as stainless steel (Somers et al., 1994; Thormar and Hilmarsson, 2010), and on heavily soiled surfaces (De Cesare et al., 2003; Brown et al., 2014) was assessed using C. jejuni NCTC 11168 biofilms formed on sterile stainless steel coupons. There was a significant reduction of crystal violet staining following DNase I treatment (**Figure 5A**). Crystal violet staining of the coupons showed no detectable biofilm following static aerobic incubation in the presence of DNase I, however significant levels of biofilm formation were observed when DNase I was not present (**Figure 5A**). In order to mimic environments where heavy soiling occurs, C. jejuni NCTC 11168 cultures were incubated statically in Brucella medium containing 5% v/v chicken juice. Chicken juice is a complex, undefined exudate obtained from defrosted whole chickens (Birk et al., 2004, 2006) and has a high protein and lipid content, and its presence results in increased biofilm formation due to its ability to condition abiotic surfaces (Brown et al., 2014). DNase I treatment of biofilms formed in the presence of 5% v/v chicken juice did result in a significant reduction of staining compared to untreated biofilms, although there was some residual staining, suggesting that on heavily soiled surfaces DNase I treatment does not provide the same level of biofilm degradation as observed in culture medium only (**Figure 5B**).

#### Biofilms Allow Genetic Transfer of Antibiotic Resistance to *C. jejuni*

Given the presence and structural importance of the eDNA we hypothesized that addition of exogenous DNA may further

(fifth bar) or fresh *C. jejuni* NCTC 11168 culture (sixth bar) and incubated for a further 48 h. The following controls were also prepared: deviation. Statistically significant results, as determined using the Mann–Whitney U test, are indicated using an asterisk (\**P* < 0.05).

increase biofilm formation. This was tested by the addition of 2µg of genomic DNA, isolated from a C. jejuni NCTC 11168 strain expressing a GFP protein and containing an antibiotic resistance marker. Addition of genomic DNA did not lead to significant differences in the levels of crystal violet staining (**Figure 6A**). This indicates that although eDNA is essential for biofilm formation and structural stability, in contrast to previous research on C. jejuni 81–176 biofilms (Svensson et al., 2009, 2014), exogenous DNA does not act synergistically with eDNA within the C. jejuni NCTC 11168 and 81116 biofilms.

While exogenous genomic DNA was not able to increase biofilm formation, genetic transfer of the antibiotic resistance marker was detected in both the planktonic and biofilmassociated cells (**Figures 6B,C**). Chloramphenicol-resistant colonies were recovered from both planktonic and biofilm phases following addition of C. jejuni NCTC 11168 cj0046::gfp−CmR genomic DNA to static cultures of the wild-type NCTC 11168 and 81116 strains. No resistance was observed in cultures not containing C. jejuni NCTC 11168 cj0046::gfp−CmR genomic DNA, suggesting that neither planktonic or biofilm cultures of strains NCTC 11168 or 81116 are naturally resistant to chloramphenicol at the levels used in these assays (10µg/ml). Where genomic DNA had been added to the suspension at the start of static incubation, resistant cells were present in both planktonic (**Figure 6B**) and biofilm (**Figure 6C**) cultures. In cultures where genomic DNA had been added at a later (24 h) time point, lower levels of chloramphenicol-resistance were observed (**Figures 6B,C**).

### Discussion

Microbial biofilms constitute an important problem for the food industry. There is an increasing body of evidence that biofilms can aid survival of C. jejuni in the food chain. C. jejuni has previously been shown to form both single (Joshua et al., 2006) and multispecies (Sanders et al., 2007) biofilms, and biofilm formation has also been demonstrated on food chain relevant materials such as stainless steel (Peyrat et al., 2008; Sanders et al., 2008; Brown et al., 2014) and in food chain relevant environmental conditions (Reuter et al., 2010; Brown et al., 2014). While the phenomenon of biofilm formation is well established for C. jejuni, there is less information available on the composition and role of the extracellular matrix in the processing environment. Biofouling of surfaces is

using the Mann–Whitney U test, are indicated using an asterisk (\**P* < 0.05).

a problem within the food industry, where organic materials are present, and areas of attention have not only been on antimicrobial treatment, but also on biofilm dispersal and prevention. Combination treatment including various enzymatic treatments, surfactants and chelating agents may provide a suitable alternative to the chemical treatments currently in use for biofilm degradation within food processing areas (Lequette et al., 2010). The use of DNase I is an example of one such enzymatic treatment.

Treatment of biofilm-based bacterial infections with DNases has increased in recent years, and the human recombinant DNase dornase alpha (Pulmozyme) is now frequently used in the treatment of cystic fibrosis (Konstan and Ratjen, 2012). DNase I is expensive to produce, and hence the use of DNase I on biofilms has been limited to medical applications, for example inner ear infections (Thornton et al., 2013) and wound biofilm control (Swartjes et al., 2013). More recently investigations have also been carried out into the activity of enzyme treatments with foodborne bacterial pathogens such as Listeria monocytogenes. For example, L. monocytogenes biofilms formed on stainless steel can be removed by both DNase I and Proteinase K treatments (Nguyen and Burrows, 2014), similar to reported here for C. jejuni biofilms.

The eDNA within the extracellular matrix appears to have multiple functions, depending on the bacterial species investigated. Previous research in P. aeruginosa biofilms has shown that eDNA can not only provide structural stability at early stages of biofilm formation (Whitchurch et al., 2002) but is also found to be localized to specific areas of the biofilm as it matures (Ma et al., 2009), again suggesting a structural role for eDNA in P. aeruginosa biofilm organization and expansion (Gloag et al., 2013), with DNase I treatment of developing biofilms leading to significant decreases in biofilm levels. DNA can be used as nutrient source by E. coli, Shewanella, and P. aeruginosa when exposed to phosphate and carbon deficient environments (Palchevskiy and Finkel, 2006; Pinchuk et al., 2008; Mulcahy et al., 2010). Since bacteria within the biofilm are typically immobilized, DNA could provide an easily obtainable food source. Finally, for naturally competent bacteria such as C. jejuni, the eDNA can contribute to the spread of genetic traits within populations, both in the biofilm and in the planktonic populations. Genetic material can be transferred within the biofilm either by direct cell to cell transmission or uptake of exogenous DNA. Conjugation within biofilms is a well reported phenomenon, with examples reported in mixed species oral biofilm models (Hannan et al., 2010), drinking water

systems (Lisle and Rose, 1995) and within bacterial populations colonizing the nasopharynx (Marks et al., 2012). Recent work has shown that C. jejuni strains NCTC 11168 and 81–176 in microaerobic cultures are able to transfer genetic material between bacterial cells both within biofilms and planktonic suspension (Bae et al., 2014; Svensson et al., 2014). The work presented here shows that C. jejuni is also able to utilize exogenously added DNA for acquisition of genetic traits. This transfer is also able to occur in aerobic conditions, more closely resembling the conditions C. jejuni encounters while in the food chain.

We demonstrate here that eDNA is an important component of the C. jejuni extracellular matrix at all stages of maturation. This is in contrast to P. aeruginosa, which become less susceptible to DNase I treatment as the biofilm matures (Whitchurch et al.,

#### FIGURE 6 | Continued

81116 biofilms were allowed to develop for 48 h in the presence of 2 µg *C. jejuni* NCTC 11168 *cj0046*::*gfp*+−CmR genomic DNA. Supplementation with eDNA did not lead to changes in biofilm formation (A). Plating both planktonic (B) and biofilm (C) cells on both Brucella media and Brucella media supplemented with 10µg/ml chloramphenicol shows emerging chloramphenicol resistant cells suggesting integration of the chloramphenicol resistance gene, via natural transformation, into the genomes of both planktonic and biofilms cells. Error bars show standard deviation.

2002). Some outer membrane and flagella proteins have been identified as been important in C. jejuni biofilm formation, but to date there has been little investigation of the extracellular matrix components themselves. C. jejuni produces a polysaccharide containing β1-3 and/or β1-4 linkages which is reactive to calcofluor white (McLennan et al., 2008), and hence further studies are required to distinguish between the roles of eDNA and other polysaccharides in C. jejuni biofilms.

Although eDNA has been shown to be present within the biofilms of many different bacteria, the mechanism of its release into the extracellular milieu is still under investigation. There are two main mechanisms of DNA release; secretion and cell lysis. Secretion of eDNA has been shown in several species, including Neisseria gonorrhoeae (Hamilton et al., 2005) and P. aeruginosa (Renelli et al., 2004). Although secretion of eDNA has been observed in some bacteria, it is widely accepted that lysis is a more common method of eDNA release (Wu and Xi, 2009). For instance, Staphylococcus aureus eDNA can be released via co-ordinated lysis of a subset of the population, controlled by quorum sensing (Mann et al., 2009). To date quorum sensing mechanisms have not been described in C. jejuni (He et al., 2008; Adler et al., 2014), and although it is possible that a yet unknown quorum sensing system controls co-ordinated eDNA release in C. jejuni, this will require further investigation. P. aeruginosa biofilms showed higher concentrations of eDNA within the biofilm when cultures were supplemented with salmon sperm DNA (Chiang et al., 2013), suggesting that some biofilm-forming bacteria are able to utilize eDNA from several sources. Our results suggest that although C. jejuni NCTC 11168 and 81116 are able to utilize exogenous DNA, this does not lead to a net increase in biofilm formation. In contrast, addition of eDNA to C. jejuni 81– 176 biofilm cultures led to increased biofilm biomass (Svensson et al., 2014).

Another problem frequently encountered within food processing environments is the presence of food product debris. This presence of this debris on surfaces can lead to surface conditioning and increased bacterial attachment, as observed with chicken juice and C. jejuni (Brown et al., 2014). The attachment of L. monocytogenes to stainless steel surfaces is enhanced by surface pre-conditioning with fish and meat emulsions (Gram et al., 2007), and surface conditioning by chicken juice has been shown to enhance C. jejuni biofilm formation (Brown et al., 2014). Surface conditioning can also decrease the effectiveness of chemical cleaning products, leading to reduced killing or biofilm degradation (Gram et al., 2007). In heavily soiled environments broad spectrum enzymatic treatments may provide a useful and effective addition to current cleaning regimes, as they are able to degrade not only biofilm extracellular matrix, but potentially also the conditioning layer. Our results show that DNase I treatment is able to significantly reduce C. jejuni biofilms formed on surfaces conditioned with chicken juice, suggesting that DNase I treatment could provide a useful addition to current treatment regimens.

It should be noted that we found DNase I treatment had no effect on cell viability, only biofilm shedding. This is as expected since DNase I is only in contact with the DNA of the extracellular matrix, reducing the structural integrity of the colonies forming the biofilm, but is not able to cause a loss of viability in bacterial cells with intact membranes. This means that although the DNase I treatment provides a rapid and effective method of biofilm dispersal it would best be used in combination with antimicrobial treatments, ensuring effective biofilm degradation and bacterial inactivation.

In conclusion, eDNA is an essential component of the C. jejuni biofilm and its degradation results in a reduction of biofilm levels below detection levels (Tresse et al., 2006). Treatment of abiotic surfaces containing C. jejuni biofilms with DNase I also prevents re-establishment of biofilms, possibly allowing more efficient antimicrobial treatment. DNase I treatment is effective on food chain relevant surfaces and hence could provide a useful addition to current food chain cleaning regimes.

## Author Contributions

HB, MR, RB, and AV designed the study. HB, KH, and MR performed the experimental work and analyzed the data. HB prepared the manuscript, and KH, MR, RB, and AV contributed to the final manuscript.

### Acknowledgments

The authors wish to thanks members of the IFR Campylobacter research group, Duncan Gaskin for the GFP-expressing C. jejuni strain, and Gary Barker for helpful discussions. We would also like to thank Maddy Houchen and Val Russell for media and laboratory support. We gratefully acknowledge the support of the Biotechnology and Biological Sciences Research Council (BBSRC) via the BBSRC Institute Strategic Programme (BB/J004529/1) and a BBSRC CASE studentship (BB/I15321/1) with CASE funding from Campden BRI.

### Supplementary Material

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fmicb. 2015.00699

#### References

**Conflict of Interest Statement:** Roy P. Betts is a full time employee of Campden BRI, which provided additional funding for the PhD-studentship of Helen L. Brown. The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2015 Brown, Hanman, Reuter, Betts and van Vliet. This is an openaccess article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Biofilm spatial organization by the emerging pathogen *Campylobacter jejuni*: comparison between NCTC 11168 and 81-176 strains under microaerobic and oxygen-enriched conditions

Hana Turonova1, 2, 3, Romain Briandet 4 , Ramila Rodrigues 1, 2, Mathieu Hernould2 , Nabil Hayek 5 , Alain Stintzi 5 , Jarmila Pazlarova3 and Odile Tresse1, 2 \*

1 SECALIM UMR1014, Institut National de la Recherche Agronomique, Nantes, France, 2 LUNAM Université, Oniris, Université de Nantes, Nantes, France, 3 Department of Biochemistry and Microbiology, Faculty of Food and Biochemical Technology, University of Chemistry and Technology, Prague, Czech Republic, 4 MICALIS UMR1319, Institut National de la Recherche Agronomique, Massy, France, 5 Department of Biochemistry, Microbiology and Immunology, Faculty of Medicine, University of Ottawa, Ottawa, ON, Canada

#### *Edited by:*

Michael Gänzle, University of Alberta, Canada

#### *Reviewed by:*

Ian F. Connerton, University of Nottingham, UK Bastien Fremaux, Institut du Porc, France

#### *\*Correspondence:*

Odile Tresse, UMR-INRA 1014 SECALIM, Oniris, Rte de Gachet, 44307 Nantes, France<EMAIL_ADDRESS>
#### *Specialty section:*

This article was submitted to Food Microbiology, a section of the journal Frontiers in Microbiology

*Received:* 03 April 2015 *Accepted:* 29 June 2015 *Published:* 13 July 2015

#### *Citation:*

Turonova H, Briandet R, Rodrigues R, Hernould M, Hayek N, Stintzi A, Pazlarova J and Tresse O (2015) Biofilm spatial organization by the emerging pathogen Campylobacter jejuni: comparison between NCTC 11168 and 81-176 strains under microaerobic and oxygen-enriched conditions. Front. Microbiol. 6:709. doi: 10.3389/fmicb.2015.00709 During the last years, Campylobacter has emerged as the leading cause of bacterial foodborne infections in developed countries. Described as an obligate microaerophile, Campylobacter has puzzled scientists by surviving a wide range of environmental oxidative stresses on foods farm to retail, and thereafter intestinal transit and oxidative damage from macrophages to cause human infection. In this study, confocal laser scanning microscopy (CLSM) was used to explore the biofilm development of two welldescribed Campylobacter jejuni strains (NCTC 11168 and 81-176) prior to or during cultivation under oxygen-enriched conditions. Quantitative and qualitative appraisal indicated that C. jejuni formed finger-like biofilm structures with an open ultrastructure for 81-176 and a multilayer-like structure for NCTC 11168 under microaerobic conditions (MAC). The presence of motile cells within the biofilm confirmed the maturation of the C. jejuni 81-176 biofilm. Acclimation of cells to oxygen-enriched conditions led to significant enhancement of biofilm formation during the early stages of the process. Exposure to these conditions during biofilm cultivation induced an even greater biofilm development for both strains, indicating that oxygen demand for biofilm formation is higher than for planktonic growth counterparts. Overexpression of cosR in the poorer biofilm-forming strain, NCTC 11168, enhanced biofilm development dramatically by promoting an open ultrastructure similar to that observed for 81-176. Consequently, the regulator CosR is likely to be a key protein in the maturation of C. jejuni biofilm, although it is not linked to oxygen stimulation. These unexpected data advocate challenging studies by reconsidering the paradigm of fastidious requirements for C. jejuni growth when various subpopulations (from quiescent to motile cells) coexist in biofilms. These findings constitute a clear example of a survival strategy used by this emerging human pathogen.

Keywords: *Campylobacter jejuni*, biofilm, CLSM, oxidative stress, CosR

### Introduction

Campylobacter has emerged as the leading cause of bacterial foodborne infections in developed countries (Epps et al., 2013; Golz et al., 2014). The resulting disease in humans, campylobacteriosis, is characterized by acute enteritis with the presence of blood and leukocytes in a stool, abdominal pain, and fever (Cameron et al., 2012; Lu et al., 2012). It is also associated with late onset complications such as Guillain-Barré syndrome, its variant Miller-Fisher syndrome (Salloway et al., 1996; Nachamkin et al., 1998; Kudirkiene et al., 2012), and inflammatory bowel diseases (Kaakoush et al., 2014). The underlying molecular mechanisms responsible for its pathogenesis, persistence, and survival seem to be unique to Campylobacter as compared to other invasive foodborne bacterial pathogens (Listeria monocytogenes, Salmonella enterica, and Staphylococcus aureus). These features might result from high level of genomic polymorphism, restricted catabolic capacity, self-regulation and deregulation of genes, and other undefined survival routes.

The main reservoir of Campylobacter is the intestinal tract of birds and other endothermic animals, especially livestock. It is primarily isolated from poultry and, to a lesser extent, pork and beef. The infection of the human host is generally caused by the consumption of undercooked and mishandled poultry or by cross-contamination of cooking tools and fresh vegetables (Butzler, 2004; Guyard-Nicodeme et al., 2013). A significant increase in the prevalence of campylobacteriosis cases has been observed over the past 5 years in the EU, based on quantitative epidemiological analyses from farms to retail outlets (EFSA, 2012, 2013). A baseline survey, conducted in 28 European countries in 2010, indicated that 71.2% of broiler batches and 75.8% of broiler carcasses were contaminated by Campylobacter (EFSA, 2010). These data were reinforced by an in-depth analysis over a 3-year period at the UK-wide level showing that in over 37 abattoirs (representing almost 90% of the total UK slaughter throughput), 79.2% of the slaughter batches were positive for Campylobacter (Lawes et al., 2012). In addition, 87.3% of the broiler carcasses were contaminated by Campylobacter with 27.3% of them showing a load over 1000 cfu.g−1 (Powell et al., 2012). In the USA, 168 pathogen-food combinations of 14 major pathogens across 12 food categories were compared (Batz et al., 2012). The combination "Campylobacter-poultry" reached the first rank in terms of annual disease burden including illnesses, hospitalizations, deaths, and costs. Overall, these exhaustive data on Campylobacter contamination indicate that this microorganism can survive outside of its reservoir through breeding farms, slaughterhouses and food processing, defying environmental conditions, and human defense mechanisms.

The main pathogenic species, Campylobacter jejuni, has been isolated in more than 80% of the campylobacteriosis cases (Moore et al., 2005). Being an obligate microaerophilic bacterium, Campylobacter has to develop adaptation strategies to survive oxidative conditions from food environments and macrophage attacks. It has been suggested that adhesion to surfaces and formation of biofilms could be one of the strategies used to maintain C. jejuni survival (Nguyen et al., 2012). Moreover, the bacterium can be sheltered in mixed species biofilms (Sanders et al., 2007; Ica et al., 2011). C. jejuni can form three different types of biofilm: (i) a structure attached to an abiotic surface, (ii) aggregates floating in the liquid culture, or (iii) a pellicle formed at the gas/liquid interface (Joshua et al., 2006). Biofilm formation occurs within 48 h of cultivation with cell detachment becoming prominent after a prolonged cultivation time (Sanders et al., 2007; Ica et al., 2011). In line with other biofilm-producing foodborne bacteria, the substratum composition and its physicochemical properties influence the biofilm formation of C. jejuni (Nguyen et al., 2011). These properties could play an important role in the early stages of biofilm formation when cells adhere to the surface. This assumption is supported by a variation in adhesion rates to inert surfaces, such as nitrocellulose membrane, glass, and stainless steel (Joshua et al., 2006; Kalmokoff et al., 2006; Gunther and Chen, 2009). The wide range of adhesion capability in Campylobacter spp. also raises the question of biological fitness among strains, in regards to their ability to attach irreversibly to a surface and initiate biofilm formation (Joshua et al., 2006; Sulaeman et al., 2010; Teh et al., 2010).

Molecular mechanisms regulating biofilm formation of C. jejuni are still poorly understood. So far, genes described to be involved in the process include those responsible for cell motility (flaA, flaB, flaC, flaG, fliA, fliS, and flhA; Joshua et al., 2006; Kalmokoff et al., 2006; Reeser et al., 2007; Reuter et al., 2010), cell surface modifications (peb4, pgp1, and waaF; Asakura et al., 2007; Naito et al., 2010; Frirdich et al., 2014), quorum sensing (luxS; Reeser et al., 2007), and stress response (ppk1, spoT, cj1556, csrA, cosR, and cprS; Candon et al., 2007; Fields and Thompson, 2008; McLennan et al., 2008; Svensson et al., 2009; Gundogdu et al., 2011; Oh and Jeon, 2014). It was found that biofilm formation is flagellum-mediated as the first step of the process—cellular adhesion—requires presence of flagella, although its functionality is not crucial for the biofilm initiation (Svensson et al., 2014). Other components essential for development of biofilm structure are extracellular DNA (eDNA) and DNA-binding protein Dps, whose presence is required for proper formation of microcolonies and structuralization of biofilm (Svensson et al., 2014; Brown et al., 2015). Genes regulating biofilm formation were not fully identified so far. Experiments using knock-out and knock-down mutants of various regulators revealed several genes influencing the process of biofilm formation. Except of aforementioned motility apparatus regulated by flhA (Kalmokoff et al., 2006), and functional quorum sensing luxS (Reeser et al., 2007), other regulators involved mostly in stress response were found to be critical for biofilm formation. Interestingly, while mutants lacking genes responsible for oxidative stress response such as cj1556 and csrA were defective in biofilm formation (Fields and Thompson, 2008; Gundogdu et al., 2011), knock-out/down of genes responsible for general stress response (spoT, ppk1, and cprS) resulted in increased biofilm formation suggesting that the process represents alternative pathway of

**Abbreviations:** BFI, Biofilm index; CLSM, Confocal laser scanning microscopy; MAC, Microaerobic conditions; OEC, Oxygen-enriched conditions; OECC, Cultivation in OEC; OECA, Acclimation to OEC; ROS, Reactive oxygen species; TCS, Two-component system.

stress defense in Campylobacter (Candon et al., 2007; McLennan et al., 2008; Svensson et al., 2009). Another regulator possibly involved in biofilm formation is gene cosR. This orphan twocomponent system (TCS) was recently discovered to be involved in the regulation pathway of ROS detoxification in C. jejuni (Hwang et al., 2011, 2012). It was previously reported that CosR regulates transcription of 93 different genes in C. jejuni (Hwang et al., 2012), it is overexpressed in sessile cells (Kalmokoff et al., 2006) and was already shown to influence biofilm formation by regulation of alkyl hydroperoxide reductase ahpC (Oh and Jeon, 2014). All these facts suggest that CosR might play significant role in biofilm formation of C. jejuni.

So far, analyses of pure cultures have mostly been carried out in an optimal growth atmosphere and were focused on the strain NCTC 11168 (Kalmokoff et al., 2006; Ica et al., 2011). Using colorimetric assessment methods (Crystal violet and Congo red assays) for biofilm detection in glass tubes, Reuter et al. (2010) showed that aerobic cultivation enhanced C. jejuni NCTC 11168 biofilm. In a previous study, we have shown that the strain 81-176, grown under controlled oxygenenriched conditions (19% O2, 10% CO2, and 71% N2), is able to overexpress membrane proteins involved in biofilm initiation and virulence process (Sulaeman et al., 2012). In this study, we compared the biofilm development of two C. jejuni strains responsible for human outbreaks (NCTC 11168 and 81- 176), and the effect of dioxygen (O2) on biofilm development. The usage of controlled atmosphere eliminated other factors possibly affecting biofilm formation. It was therefore possible to explore whether the increase of biofilm formation in aerobic conditions could be attributed solely to the level of oxygen and if the trend of enhanced biofilm formation is present in other strain of C. jejuni. This was evaluated, for the first time, using specific biofilm parameters (maximum height, biomass volume, and ultrastructure) from confocal laser scanning microscopy (CLSM) analyses. This non-invasive sensitive technique has been used previously to examine Campylobacter cell morphology and viability (Chantarapanont et al., 2003; Lee et al., 2004; Jang et al., 2007; Ica et al., 2011) and bacterial interactions with live tissues (Mooney et al., 2003). The CLSM has also been used for the detection of C. jejuni in mixed species biofilms (Sanders et al., 2007; Ica et al., 2011). In the present work, the impact of pretreatment and cultivation of cells in oxygenenriched conditions (OEC) on C. jejuni biofilm formation and its ultrastructural organization was investigated in comparison with cells cultivated in microaerobic conditions (MAC). In addition, analyses using an overexpressing cosR transformant were performed to determine the role of this regulator in C. jejuni biofilm development.

#### Results

#### Biofilm Development and Architecture

Two C. jejuni strains, NCTC 11168 and 81-176, were chosen in order to explore their biofilm formation capacities using CLSM with Syto 9 staining. This cell-permeable dye emits fluorescence after binding to nucleic acids and therefore allows the visualization of cells and any extracellular DNA present in the biofilm matrix. Both strains were able to form biofilm within 24 h of cultivation (**Figure 1**). At the initial stages of biofilm formation, cells gathered in clusters partially attached to the surface, forming finger-like structures. After 48 h, most of the biofilm mass remained attached to the bottom of the well. The biofilm structure evolved during the time of cultivation, increasing in both maximum height and biomass volume for both strains. However, 81-176 formed more biofilm than the NCTC 11168 strain (after 48 h: 233.33 ± 64.63 and 130.67 ± 14.70µm, respectively, for the maximum height; 42.3 × 105 ± 5.7 × 105 and 0.4 × 105 ± 0.09 × 105 µm3 , respectively, for the biomass volume; n = 3). In addition, unlike NCTC 11168, the biofilm of the 81-176 strain exhibited a pronounced open ultrastructure full of voids and channels, even after 96 h of incubation (data not shown). As growth rates of both strains were similar (µmax = 0.69 h −1 for NCTC 11168 and µmax = 0.67 h −1 for 81-176), these differences in biofilm formation cannot be explained by different growth abilities. During the experiment, no formation of pellicle or floating aggregates was observed probably due to the cultivation in static conditions.

#### Cell Motility in Biofilm

Motile cells, tracked using CLSM, were observed around or inside the biofilm structure after 48 h of cultivation (Supplementary Videos). However, the motility of cells differed according to their position in the biofilm structure. The highest number of motile cells was detected at the bottom of the well (Supplementary Videos 1, 3) moving more or less freely through the structure, while the motility and the number of motile cells decreased in the middle part of the biofilm (Supplementary Videos 2, 4). Furthermore, high number of motile cells was detected within the biofilm structure of 81-176 (Supplementary Videos 1, 2), whereas for NCTC 11168 the motile cells were detected mostly outside the biofilm (Supplementary Videos 3, 4).

#### Effect of Oxygen on Biofilm Formation of NCTC 11168 and 81-176 *C. jejuni* Strains

Two different approaches were used to evaluate the effect of subinhibitory oxygen concentration on biofilm formation of

FIGURE 1 | *C. jejuni* NCTC 11168 and 81-176 biofilm architecture and development are different after incubation for 24 and 48 h in MAC (5% O2 , 10% CO2 , 85% N2 ). The CLSM images represent an aerial view of biofilm structures with the shadow projection at the bottom. The structures were visualized using Syto 9, an intercalating agent staining the nucleic acids.

two strains with different biofilm forming ability (NCTC 11168 and 81-176). Firstly, biofilms were cultivated under controlled oxygen-enriched conditions (OECc) as described previously by Sulaeman et al. (2012). In OEC, the same concentration of CO2 (10%) as in MAC was maintained, while the O2 concentration was increased to a sublethal level (19% O2 in OEC vs. 5% in MAC). This enabled the evaluation of the effect of increased O2 concentration on biofilm development of C. jejuni regardless of its capnophilic nature requiring increased concentration of CO2. Biofilm volume of both strains was significantly increased (P < 0.01) when cultivated in OECc (**Figure 2** and Supplementary Table 1). Incubation time and O2 concentration had a significant effect (P < 0.01) on increased biomass production in OECc when compared to MACc. Interestingly, some significant differences in both maximum height and biomass volume (P < 0.01) remained between the two strains even after cultivation in OEC, with a greater biofilm development for 81-176 than for NCTC 11168, indicating that strain biology impacts biofilm formation (Supplementary Table 1). This was confirmed by formation of a denser compact biomass for NCTC 11168 biofilm while 81- 176 induced more voids and open water channels across the biofilm.

In the second approach, both strains were acclimatized to OEC (OECa) prior to biofilm formation in MAC. Acclimatized cells of both strains formed significantly larger biofilms than nonacclimatized ones after 24 h of cultivation, as expressed by the fold changes in maximum height and biomass volume values (**Figure 3A**). Conversely, the acclimatization of cells to OEC was no longer an advantage for biofilm formation after 48 h, as demonstrated by reduction of biofilm formation for both strains. This was also confirmed by statistical analyses, with the highest F-ratios of the interaction effect between "Incubation time" and "O2 pretreatment," showing higher variation in maximum height and biomass volume, than for the other factors (Supplementary Table 2).

In order to distinguish the effect of OEC prior to or during C. jejuni biofilm formation, biofilm development was compared between the cells acclimatized to OEC and the cells subjected to OEC during biofilm formation (OECa and OECc, respectively; **Figure 3B**). Although the fold change (OECc/OECa) was not in favor of NCTC 11168 biofilm formation during the first 24 h, after 48 h both strains cultivated in OECc showed enhanced biofilm formation with a marked difference in biomass volume for 81-176. This was confirmed statistically with a significant effect of OEC treatment (P < 0.0001) for O2 treatment, and the interaction between "Incubation time" and "O2 treatment" with the highest F-ratios (Supplementary Table 3).

#### Role of *cosR* in Biofilm Development

A second copy of C. jejuni gene cosR and its promoter were inserted into the poorer biofilm-forming strain NCTC 11168 to determine its role in C. jejuni biofilm formation. This construction, with an ectopic copy of the cosR gene and its promoter, enabled to double the expression of the transcript level of cosR in the cells (Supplementary Figure 1) in a same manner as in the cosR-overexpressing strain obtained by Hwang et al. (2011) and used by Oh and Jeon (2014). Then, the parental NCTC 11168 strain and the cosR overexpressing strain, namely transformant (TrfcosR), were compared for their ability to adhere to an inert surface and to develop a biofilm (**Figure 4**). Using the BioFilm Control Ring Test R , a significantly higher 1BFI was obtained (P = 0.0007) for the transformed strain, indicating its greater ability to adhere to inert surfaces (**Figure 4A**). In addition, using the crystal violet assay, the transformant showed enhanced biofilm formation after 24 and 48 h (P = 0.0006 and 0.02, respectively) but not after 72 h (P > 0.05) when compared with its parental strain (**Figure 4B**). The CLSM observations and biofilm analyses indicated that the transformant formed significantly more (P < 0.01) biofilm than its parental strain (**Figure 5**, Supplementary Table 4). In addition, the maximum height and biomass volume reached by the transformant was not significantly different from those obtained with the strongest biofilm-forming strain 81-176 (Supplementary Table 5). These data showed that the presence of two genes encoding cosR significantly enhanced biofilm development in MAC (592.7-times higher biomass volume after 24 h). Interestingly, this was correlated with the formation of an open biofilm ultrastructure with voids and water channels similar to the one described for 81-176 (**Figures 1**, **5A**). Comparison of genomic sequences using xBASE2 (Chaudhuri et al., 2008) showed that the cosR gene (cj0335c and cjj0379c, respectively) and its flanking regions are 100% identical in NCTC 11168 and 81-176. Both strains carry the exact same form of the gene. Therefore, some other mechanisms, related to the cosR sequence and its flanking regions, for regulating the C. jejuni biofilm formation, should exist. Moreover, unlike the two wild strains, an increased O2 concentration during cultivation did not promote biofilm formation of the transformant (**Figure 5**). These data indicate that a second ectopic copy of cosR enhanced biofilm development by promoting a complex architecture of C. jejuni biofilm irrespective of O2 demand. Nevertheless, further experiments should be performed to evaluate cosR transcript level and CosR expression throughout all phases of biofilm development.

#### Discussion

As the leading cause of bacterial foodborne diseases, whose incidence has been significantly increasing during the recent years in Europe (EFSA, 2010, 2012, 2013), this pathogen has to adapt and survive environmental conditions outside and inside its main hosts, particularly oxidative stress. In this study, we have shown that C. jejuni can form biofilm in static conditions with a clearly defined finger-like structure. Our observation is consistent with previous studies indicating that C. jejuni could develop monospecies biofilms (Kalmokoff et al., 2006; Asakura et al., 2007; Corcoran and Moran, 2007; Reeser et al., 2007; Fields and Thompson, 2008; Hanning et al., 2008; McLennan et al., 2008; Sanders et al., 2008; Gunther and Chen, 2009). Both examined strains were able to produce a biofilm, although their maximum height, biomass volume, and ultrastructure differed significantly between the two strains. In previous studies, stronger adhesion to an inert surface was observed for 81-176 than for NCTC 11168 (Gunther and Chen, 2009; Sulaeman et al., 2010; Teh et al., 2010). Although the adhesion strength could not be fully correlated to the capability of bacterial species to form biofilms, biofilm initiation is crucial to anchor the embryonic core of the biofilm. Our qualitative and quantitative data indicated that

FIGURE 4 | The NCTC 11168 *cosR* overexpressing transformant enhanced cell adhesion to inert surface and biofilm formation in comparison with its parental strain. Adhesion to an inert surface (A) and biofilm formation (B) of C. jejuni NCTC 11168 (white bars) and the cosR overexpressing transformant (TrfcosR) (black bars) strains. Bacterial adhesion was determined after 2 h by calculating the BioFilm Index (BFI) using the

BioFilm Ring Test®. Biofilm formation was measured in 24-well microtitre plates at 24, 48, and 72 h using the crystal violet assay. Error bars represent the standard deviation of three independent experiments. Asterisks indicate significant differences (P < 0.05) between the parental strain and the transformant. A dashed line represents detection limit of the crystal violet assay.

FIGURE 5 | The CosR is responsible for biofilm maturation in *C. jejuni*. Biofilm structure of C. jejuni NCTC 11168 and the cosR overexpressing transformant (TrfcosR) after incubation for 24 and 48 h in MAC (black bars) or OEC (white bars). (A) The CLSM images representing an

NCTC 11168 formed a thin but compact multilayered biofilm without achieving a more complex organization during the time of incubation. In contrast, the 81-176 strain was able to form a thick biofilm with an open ultrastructure composed of voids and channels. This kind of heterogeneous structure is considered to be the signature of a mature biofilm. It enhances the formation of convective flows bringing nutrients to cell aggregates and draining metabolic waste from cells in these aggregates (Donlan and Costerton, 2002). The heterogeneity of the 81-176 biofilm was confirmed by tracking the motile cells within the C. jejuni 81-176 biofilm. In contrast to many other bacteria, C. jejuni is able to maintain the expression level of genes responsible for cell motility and flagella biosynthesis when grown in biofilms (Joshua aerial view of biofilm structures with the shadow projection on the right. (B) TrfcosR biofilm development in comparison to the parental strain expressed as a fold changes of maximum height and biomass volume. Statistical data are presented in Supplementary Tables 4, 5.

et al., 2006; Kalmokoff et al., 2006; Asakura et al., 2007; Reeser et al., 2007). In our study, we observed the presence of motile, less motile and sessile cells, indicating that the biofilm is composed of cells in different physiological states. Due to the biofilm organization, different cell phenotypes coexist in the structure and therefore a wide range of cells can be found in the biofilm, from dormant to motile cells. As in nature (ex vivo or in vivo) C. jejuni cells may encompass various physiological states, biofilm could be considered as a model of mixed subpopulations of C. jejuni which could be found in food products, food-processing plants, in poultry gut, or human digestive tract.

Although C. jejuni is sensitive to increased concentrations of oxygen, absence of oxygen in anaerobic conditions induces cell death. C. jejuni requires a basal amount of available oxygen to maintain the processes essential for respiration and multiplication (Kelly, 2008). The availability of dissolved oxygen is therefore one of the main environmental parameters for the survival of C. jejuni. Previous studies showed that aerobic conditions enhanced biofilm formation of the strain NCTC 11168 (Asakura et al., 2007; Reuter et al., 2010). In those studies, biofilms grown in glass tubes or in 24 well plates were detected by crystal violet or Congo red after exposition to air and air supplemented with CO2. These colorimetric assays showed enhancement of biofilm formation under the oxidative stress, but could not predict whether and how the biofilm structure would change. The controlled O2 gaseous conditions, respecting the capnophilic nature of C. jejuni, and the use of CLSM allowed us not only to quantify the amount of biofilm, but also to evaluate any structural changes caused by increased oxygen concentration. In accordance to our expectations, we did observe an increased biofilm formation for both strains under OEC. Moreover, the data obtained using CLSM suggest that the response to an increased O2 level is strain-dependent. Although the biofilm formation for both strains was enhanced, the ultrastructures were remarkably different. The poorer biofilm forming NCTC 11168 produced more voluminous biofilm without increasing its thickness and without switching to a maturation phase as observed for 81-176. As the physiological state of cells may correspond to their close environment, the cell response to environmental conditions could differ according to its location in the biofilm structure. The formation of voluminous flat biofilm may be beneficial for NCTC 11168 under OEC, as a smaller area, and therefore a smaller number of cells, is exposed to the malignant effect of oxygen. On the other hand, the 81- 176 strain increased in both biofilm volume and height, keeping the porous ultrastructure of the biofilms produced under MAC. It seems like the strain disregards the negative effects of an increased oxygen level and is supported to multiply and form a mature biofilm composed of mixed subpopulations of cells. The biofilm organization may therefore offer a favorable oxygen tuning niche for C. jejuni. These findings indicate that oxygen growth requirements of C. jejuni are not as fastidious when cells are organized in biofilm. Consequently, the paradigm of fastidious requirements for C. jejuni growth (Jones, 2001; Park, 2002) should be reconsidered according to the cell physiological state and cell population cooperation.

Unlike well-studied aerobes, C. jejuni lacks specific and global regulators involved in oxidative stress resistance, such as SoxRS, OxyR, or RpoS (Garenaux et al., 2008). C. jejuni carries two Fur homologs, Fur, and PerR (peroxide stress regulator), which regulate iron homeostasis and contribute to the oxidative stress response (Van Vliet et al., 2002). Recently, Hwang et al. (2011, 2012) have suggested that the orphan TCS Cj0355c could be involved in the oxidative stress response and named it CosR. The protein product of cosR shares 60% amino acid identity with Hp1043, a TCS response regulator element from the close relative Helicobacter pylori. Deletion of hp1043 induced death of H. pylori in the same way as it has been observed for cosR and C. jejuni (Stahl and Stintzi, 2011). However, the hp1043 gene has been successfully substituted by C. jejuni cosR (Muller et al., 2007) suggesting that CosR exhibits similar biological functions to Hp1043.

In this study, the essential gene cosR was overexpressed in the poorer biofilm-forming strain, NCTC 11168, in order to investigate the role of this TCS in biofilm formation and structuring. In our study, the adhesion of cells to inert surfaces was correlated to the biofilm formation detected by Crystal violet and analyzed by CLSM. All three different detection techniques led to the same conclusion. The significantly greater adhesion to an inert surface and the increased biofilm formation of the transformant (TrfcosR) revealed that the expression of this gene is connected to biofilm formation. This was also confirmed by the CLSM experiments, which showed a much greater thickness and volume of the transformant's biofilm under MAC than those observed for the parental strain. This result is in accordance with the previously published work describing increased expression of CosR in C. jejuni NCTC 11168 biofilm as compared to planktonic counterparts (Kalmokoff et al., 2006), although Oh and Jeon (2014) observed decrease of biofilm formation in strain overexpressing cosR. This discrepancy might be explained by looking at the structure of biofilms of parental strain and the transformant. Interestingly, the ultrastructure of the TrfcosR biofilm was found to be more similar to the one described for 81-176 than the parental strain, showing an open organization with pores and channels across the structure. Unexpectedly, when the transformant was cultivated under OEC, the maximum height and biomass volume of the biofilm were not higher than when the biofilm was produced under MAC. Nevertheless, the values still remained higher than those of the parental strain. These data indicate that cells overexpressing cosR were not stimulated by the higher O2 concentrations to enhance the biofilm formation. Thus, CosR seems to be crucial for initiation of the maturation phase of C. jejuni biofilm development. This might be the reason of arisen discrepancy between our work and the one published by Oh and Jeon (2014). The authors used Mueller Hinton broth and higher temperature of cultivation. These factors were previously found to increase the biofilm formation of C. jejuni (Reeser et al., 2007). The usage of supportive cultivation conditions in combination with enhanced initiation of biofilm maturation caused by overexpression of CosR might result in earlier dispersal of cells and microcolonies from mature biofilm. Such acceleration of dispersal would result in reduction of biofilm mass attached to the surface of the well and cosR transformant would therefore seem to be less biofilm forming. This is in accordance with our observation of the dramatic biovolume decrease after 48 h of cultivation (from 600 to 16-times more biofilm mass than the parental strain) observed for the transformant. Nevertheless, further experiments should be performed in order to confirm or refuse this hypothesis.

The regulator CosR was initially identified as a potential regulator of ROS scavengers by promoting or repressing genes encoding KatA, AhpC, and SodB in C. jejuni (Hwang et al., 2011, 2012). It was also differently expressed after a superoxide stress induced by paraquat (Garenaux et al., 2008). Binding to the promoter of luxS, CosR might also be contributing to a quorum sensing system (Hwang et al., 2011). Recently, it was also demonstrated that CosR is involved in the expression of the

antibiotic efflux pump CmeABC in C. jejuni (Hwang et al., 2012). In this study, role of CosR in the maturation of C. jejuni biofilm, independently of oxidative stress, adds a new element in favor of its pleiotropic function in the main metabolic processes allowing the survival of C. jejuni in response to environmental stresses.

In contrast to its highly restricted catabolic capacity, C. jejuni is able to develop strategies to survive environmental oxidative stress using O2 as an advantage for biofilm development. As C. jejuni is equipped to withstand oxidative stress through cooperation of subpopulations within a biofilm, further analyses are required to assess if this feature could explain the survival of this emerging pathogen in slaughterhouses, after evisceration, during food processing, or during macrophage attack. In addition, these findings advocate further studies to analyze quiescent, dormant and sessile C. jejuni cells and cell cooperation in response to environmental stresses, to identify the underlying cellular and molecular mechanisms supporting the persistence and resistance of this mysterious pathogen.

#### Materials and Methods

#### Bacterial Strains and Culture Conditions

All experiments were performed using three strains of C. jejuni: two well-documented clinical isolates 81-176 and NCTC 11168 purchased from general collections, and a cosR overexpressing transformant built for this study as described below. All strains were subcultured from the stock stored at −80◦C by cultivation on Karmali agar plates (Oxoid, UK) at 42◦C for 48 h in MAC (5% O2, 10% CO2, and 85% N2, namely MAC). Grown colonies were inoculated onto Karmali agar plates and incubated either for 24 h at 42◦C in MAC, or for 42 h in oxygen-enriched conditions (19% O2, 10% CO2, and 71% N2, namely OEC) to allow acclimation of cells to oxidative stress. The OEC were previously described as a sublethal atmosphere not repressing growth of C. jejuni NCTC 11168 and 81-176 (Sulaeman et al., 2012). The gas conditions were maintained using hermetic stainless steel jars vacuum flushed and then filled with commercially purchased gas mixture. The process was repeated two times to minimize air residua in the cultivation atmospheres. The growth rates of all tested strains were determined from cultivation in BHI (Merck, Germany) in MAC using plate counts in triplicates with the appropriated decimal dilution.

#### Construction of the *cosR* Overexpressing Strain

For the construction of the cosR overexpressing strain, the cj0355c (cosR) gene was amplified from the strain NCTC 11168 using PCR primers Cj0355c F and Cj0355c R (Supplementary Table 6). The positions of the forward and reverse primers were chosen upstream and downstream of cosR within the folB (start position at 325186) and fdxB (end position at 323902) genes, respectively, to ensure that cosR was under the control of its own promoter. The PCR product was purified using the Qiagen PCR purification kit (Toronto, ON, Canada) and then cloned into the pRRK-1 plasmid (Reid et al., 2008). The cloning step was achieved using the Clontech In-Fusion™ PCR cloning kit (Mountain View, CA, USA). Briefly, the primers were designed with 15 bp extensions that allow recombination with the nucleotides flanking the XbaI restriction site on the pRRK vector. The recombinant pRRK + cosR plasmid was transformed into Fusion-Blue competent cells and positive transformants were selected on LB agar plates supplemented with Km. The cloned plasmid with the cosR gene was extracted from the grown transformants, purified, and sequenced to confirm the absence of point mutations. The plasmid was then naturally transformed into C. jejuni NCTC 11168 grown to mid-log phase. Following incorporation of the cosR into the chromosome was achieved by heterologous recombination. The location of the inserted gene was determined by amplifying three possible insertion sites on the chromosome using the ak233, ak234, ak235, and AR56 primers (Supplementary Table 6). The expected PCR product size was detected using the ak234 and AR56 primers indicating that cosR was inserted downstream of cj0431. The NCTC 11168 + cosR + KmR strain is henceforth referred to as the cosR overexpression transformant or, for simplicity, the "transformant" or "TrfcosR." The growth rates of the parental NCTC11168 strain and the transformant were similar (µmax = 0.69 and 0.72 h−1 , respectively). The overexpression of cosR was validated using quantitative RT-PCR after RNA extraction according to Sulaeman et al. (2012) with the following modifications. The quantity of total RNA was assessed using a Nanodrop 2000 (Thermo Fisher Scientific, Courtaboeuf, France), and the integrity of the RNA was verified with an Experion™ Automated Electrophoresis Station (BioRad) using the Experion RNA StdSens Analysis Kit (BioRad) according to the manufacturer's guidelines. Absence of DNA in the samples was confirmed by PCR with primers targeting flaA (Supplementary Table 6). Only high quality RNA samples without DNA contamination were used in qRT-PCR assays.

#### Adhesion to an Inert Surface

The adhesion capability of C. jejuni strains was determined using the BioFilm Ring Test R (BioFilm Control, France) as described previously by Sulaeman et al. (2010). Briefly, each culture was pelleted and resuspended in buffered peptone water, the OD600nm was adjusted to 1 and suspension was used for inoculation of plate wells. After 2 h of cultivation under MAC at 42◦C, adhesion was determined by measuring BFI (Biofilm Index) using the BioFilm Control developed software. The BFI correlates to the number of magnetic microbeads detected after well magnetization. The 1BFI was calculated by subtracting the BFI of blank control from the BFI of the sample. The assay was repeated three times with three technical replicates for each independent culture.

#### Biofilm Formation

The crystal violet biofilm assay was used to determine the amount of biofilm produced by C. jejuni. The protocol was adapted from that described by Djordjevic et al. (2002). Briefly, 2 ml of C. jejuni suspension was inoculated in 24-well sterile microtitre plates. Each plate was incubated statically for 24, 48, or 72 h at 42◦C in MAC. After cultivation, planktonic cells were washed out and biofilm was stained with 1% crystal violet solution. The crystal violet bound to the biofilm was then eluted using 99% ethanol and the absorbance of the eluate was measured at 595 nm.

Qualitative (ultrastructural) and quantitative data (maximum thickness and biomass volume) of C. jejuni biofilm were measured on biofilm produced in 96-well polystyrene microtitre plates with a µ clear R base (thickness of 190 ± 5µm; Greiner Bio-one, Germany). Prior to the inoculation of microtitre plates, grown cells were transferred from Karmali plates into BHI, washed once and resuspended in sterile BHI to final OD600nm = 0.8 ± 0.05. The suspension was then loaded onto the microtitre plate in triplicates for each strain (250 µl per well). The plates were incubated in MAC or OEC for 4–5 h at 37◦C allowing C. jejuni cells to adhere to the substratum. After that, the bacterial suspension was carefully replaced with 250µl of sterile BHI and microtitre plates were incubated for the next 24 and 48 h at 37◦C in MAC or OEC, depending on the experiment. The µ clear R base material allows diffusion of gas molecules into the liquid media and therefore ensures formation of biofilm that is attached to the bottom of the well and not floating on the air-liquid interphase. Finally, wells containing biofilm were stained using 50µl of Syto 9 solution (Invitrogen, USA) diluted in BHI to the final concentration of 2µl/ml. The Syto 9 is cellpermeable dye intercalating with DNA and therefore staining the cells and the eDNA of biofilm matrix. All biofilms were observed using confocal laser scanning microscope (CLSM) as described below. For each condition, three independent replicates were analyzed.

#### Confocal Laser Scanning Microscopy (CLSM) Image Acquisition

Each well of the microtitre plates was scanned using the inverted Leica SP2 AOBS confocal laser scanning microscope (LEICA Microsystems, Germany) at 400 Hz with a 40x/0.8 water immersion objective lens Leica HCX Apo. The fluorophore Syto 9 was excited with a 488-nm argon laser. The whole well area was inspected to verify the presence of biofilm, then the most representative place was scanned providing a stack of horizontal planar images (512 × 512 pixels representing an area of 375 × 375µm) with a z-step of 1µm. At least one stack of horizontal planar images was acquired for each replicate.

#### Image Analysis

The stacks obtained from the microscopic observations were processed using Imaris 7.6.4 software (Bitplane, Switzerland). Images representing an aerial view of biofilm structure were rendered using the Easy 3D view with the auto-adjustment function to correct pixel intensities. Numerical data and 3D models of the biofilm structures were generated using the surface generator function of the Measurement Pro module with the minimal threshold set at 40 for the green channel (Syto 9). Only objects bigger than 10 voxels were included in the analysis. Biofilm development was normalized according to

#### References

Asakura, H., Yamasaki, M., Yamamoto, S., and Igimi, S. (2007). Deletion of peb4 gene impairs cell adhesion and biofilm formation in Campylobacter height (thickness determined from z-stacks as the last image showing consecutive signal from biofilm structures) and biomass volume (cell abundance).

#### Statistical Analyses

The numerical data obtained from Imaris were processed with STATGRAPHICS Centurion 16.1.11 software (StatPoint, Inc., Herndon, VA, USA) with the maximum height (biofilm thickness) and the biomass volume (cell abundance) as explanatory values. For all variance analyses, ANOVAs were performed to determine the individual effect of each factor and potential interacting effects with the confirmation of a normal distribution for each data set.

Assay variations were excluded from interacting effects, as they were not significantly different at the first order. The significance level was selected at 99%, consequently an effect was considered significant if its P-value was lower than 0.01. All Fratios were based on the average residual squared error. When the transformant (TrfcosR) was used, a multiple comparison using the Scheffé method was implemented in ANOVAs to classify the significant variations (at 95% confidence) according to the strains.

For cell adhesion to an inert surface and crystal violet biofilm assays, significant differences were determined using two-sided Student's t-test comparisons at a 95% significance level with the confirmation of a normal distribution for each data set.

## Author Contributions

HT, RB, and OT conceived and designed the study. NH, MH, and AS built and validated the transformant. HT, RR, and RB performed the experimental work. HT prepared the manuscript and RB and OT contributed to the final manuscript.

#### Acknowledgments

The work was financially supported by Secalim (INRA/Oniris, Nantes, France), the Czech Science Foundation Grant No. 14- 23597S, LD 14097-COST-CZ, 7AMB15FR012 Project Barrande, and the Cost BacFoodNet FA1202. We would like to thank the Micalis Unit (INRA Jouy-en-Josas, France) for material support and advice with special thanks to Julien Deschamps, who helped us to acquire the CLSM images.

#### Supplementary Material

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fmicb. 2015.00709

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**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2015 Turonova, Briandet, Rodrigues, Hernould, Hayek, Stintzi, Pazlarova and Tresse. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Adhesion, Biofilm Formation, and Genomic Features of Campylobacter jejuni Bf, an Atypical Strain Able to Grow under Aerobic Conditions

Vicky Bronnec1 , Hana Turonová ˇ 2 , Agnès Bouju1 , Stéphane Cruveiller3 , Ramila Rodrigues1 , Katerina Demnerova2 , Odile Tresse1 , Nabila Haddad1 and Monique Zagorec1 \*

1 UMR 1014 SECALIM, Oniris, Nantes, France, 2 Institute of Chemical Technology, Faculty of Food and Biochemical Technology, Department of Biochemistry and Microbiology, Prague, Czech Republic, 3 CNRS-UMR 8030 and Commissariat à l'Energie Atomique et aux Energies Alternatives CEA/DRF/IG/Genoscope LABGeM, Evry, France

#### Edited by:

Avelino Alvarez-Ordóñez, Teagasc Food Research Centre, Ireland

#### Reviewed by:

Helen Louise Brown, Cardiff Metropolitan University, UK Gary Dykes, Curtin University, Australia

\*Correspondence: Monique Zagorec<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Food Microbiology, a section of the journal Frontiers in Microbiology

Received: 21 March 2016 Accepted: 13 June 2016 Published: 30 June 2016

#### Citation:

Bronnec V, Turonová H, Bouju A, ˇ Cruveiller S, Rodrigues R, Demnerova K, Tresse O, Haddad N and Zagorec M (2016) Adhesion, Biofilm Formation, and Genomic Features of Campylobacter jejuni Bf, an Atypical Strain Able to Grow under Aerobic Conditions. Front. Microbiol. 7:1002. doi: 10.3389/fmicb.2016.01002 Campylobacter jejuni is the leading cause of bacterial enteritis in Europe. Human campylobacteriosis cases are frequently associated to the consumption of contaminated poultry meat. To survive under environmental conditions encountered along the food chain, i.e., from poultry digestive tract its natural reservoir to the consumer's plate, this pathogen has developed adaptation mechanisms. Among those, biofilm lifestyle has been suggested as a strategy to survive in the food environment and under atmospheric conditions. Recently, the clinical isolate C. jejuni Bf has been shown to survive and grow under aerobic conditions, a property that may help this strain to better survive along the food chain. The aim of this study was to evaluate the adhesion capacity of C. jejuni Bf and its ability to develop a biofilm. C. jejuni Bf can adhere to abiotic surfaces and to human epithelial cells, and can develop biofilm under both microaerobiosis and aerobiosis. These two conditions have no influence on this strain, unlike results obtained with the reference strain C. jejuni 81-176, which harbors only planktonic cells under aerobic conditions. Compared to 81-176, the biofilm of C. jejuni Bf is more homogenous and cell motility at the bottom of biofilm was not modified whatever the atmosphere used. C. jejuni Bf whole genome sequence did not reveal any gene unique to this strain, suggesting that its unusual property does not result from acquisition of new genetic material. Nevertheless some genetic particularities seem to be shared only between Bf and few others strains. Among the main features of C. jejuni Bf genome we noticed (i) a complete type VI secretion system important in pathogenicity and environmental adaptation; (ii) a mutation in the oorD gene involved in oxygen metabolism; and (iii) the presence of an uncommon insertion of a 72 amino acid coding sequence upstream from dnaK, which is involved in stress resistance. Therefore, the atypical behavior of this strain under aerobic atmosphere may result from the combination of insertions and mutations. In addition, the comparison of mRNA transcript levels of several genes targeted through genome analysis suggests the modification of regulatory processes in this strain.

Keywords: food borne pathogen, biofilm, confocal microscopy, oxidative stress, genome sequence

## INTRODUCTION

fmicb-07-01002 June 28, 2016 Time: 19:12 # 2

Campylobacter is a Gram-negative bacterium, spiral-shaped and motile. This human pathogen lives as commensal of the gastrointestinal tract of most warm-blooded animals, especially poultry but also mammals (Park, 2002). Human infection by Campylobacter is commonly associated to the consumption of contaminated poultry meat. The genus Campylobacter includes very heterogeneous species that are present in a variety of environments but more than 80% of confirmed cases of campylobacteriosis were reported to be associated to Campylobacter jejuni (EFSA and ECDC, 2016).

The clinical manifestation of campylobacteriosis is severe gastro enteritis. However, Campylobacter infection is occasionally a precursor of serious post-infectious illness, including immunereactive complications such as Guillain Barré and Miller Fisher Syndromes, two chronic and potentially fatal forms of paralysis (WHO, 2013). Since 2005, Campylobacter has been the most commonly reported human gastrointestinal bacterial pathogen in the European Union (EFSA and ECDC, 2016). In 2014, 236,851 cases of human campylobacteriosis were reported in EU. This zoonosis represents an incidence rate of 71 per 100,000 population exceeding the number of salmonellosis, which has a notification rate of 23.4 cases per 100,000 population. In addition, the cost of campylobacteriosis to public health systems and the loss of individual health and productivity were evaluated around 2.4 billion Euros per year in Europe (EFSA and ECDC, 2016) and between 1.2 and 4 billion \$ for the US (Eberle and Kiess, 2012; Batz et al., 2014). The need for controlling this pathogen along the food chain explains the numerous studies reported in the literature that aimed at understanding its metabolism and virulence.

Campylobacter jejuni presents specific growth requirements, as it is thermotolerant with an optimal growth temperature of 40–42◦C, microaerophilic (optimal O2 concentration of 5%), and capnophilic requiring 10% CO2 for an optimal growth. However, C. jejuni is able to persist in different environmental stress conditions explaining its high prevalence around the world. This food-borne pathogen has indeed developed adaptation mechanisms to survive under various harsh conditions it can encounter, from poultry gastrointestinal tract to the consumer's plate. One of the most important characteristics of this bacterium is its ability to survive in aerobic environments despite its microaerophilic nature. This suggests an ability to cope with oxidative stress mediated by environmental oxygen tension and reactive oxygen species. To survive against such stresses, biofilm formation has been suggested to be one of the strategies used by this pathogen to persist in the environment (Buswell et al., 1998; Nguyen et al., 2012; Turonova et al., 2015). Commonly, biofilms are defined as multicellular layers of bacteria embedded within a matrix of extracellular polymeric substances (EPSs; Costerton, 1995; Costerton et al., 1995; Donlan, 2002; Donlan and Costerton, 2002). C. jejuni strains have been reported to be able to form different types of biofilm characterized as a structure attached to a surface, a pellicle formed at the surface of the liquid, or aggregates floating in the liquid culture (Joshua et al., 2006). Recently, we have reported the atypical property of C. jejuni Bf, a strain able to grow on plates under aerobic atmosphere, thus with a very low concentration of CO2 (0.035%), but with 21% O2 (Rodrigues et al., 2015). The possible growth of C. jejuni strains under aerobiosis and after various oxidative stresses was previously reported (Chynoweth et al., 1998; Garénaux et al., 2008b; Hinton, 2016). The aim of this study was to investigate the ability of C. jejuni Bf to adhere to biotic and abiotic surfaces and to form biofilm. We compared the behavior of this strain under both microaerobiosis and aerobiosis to determine a possible increased capacity to resist to the presence of high level of O2, which can be encountered during meat products processing and storage. Finally, genome comparison was also performed in order to detect genetic elements putatively involved in the phenotype of this strain. For that purpose, the draft genome (Bronnec et al., 2016) was completed and the gene and metabolic repertoires of C. jejuni Bf were compared to those of other complete or draft genomes.

## MATERIALS AND METHODS

#### Bacterial Strain and Culture Conditions

Stains used in this study are presented **Table 1**. C. jejuni strains were stored at −80◦C in Brain Heart Infusion broth (BHI) containing 20% (vol/vol) glycerol. Prior to each experiment frozen cells were streaked on Karmali agar plates (Oxoid Limited, UK), incubated at 42◦C for 24 h under microaerobic conditions in CampyGen sachet (Oxoid Limited, UK): 5% oxygen, 10% carbon dioxide, and 85% nitrogen.

As described previously by Rodrigues et al. (2015), C. jejuni Bf cells can be acclimated to aerobic conditions (namely AAC cells for aerobically acclimated cells). This was performed by sub-culturing three times (once for 48 h and then twice 24 h) on Karmali agar plates under aerobiosis (air; Rodrigues et al., 2015). In order to maintain the same conditions for all samples, cultures under microaerobiosis were identically performed three times under microaerobiosis (MAC cells for microaerobic conditions).

## Adhesion to Inert Surfaces

The adhesion capability was evaluated using BioFilm Ring Test R (BioFilm Control, France) as described by Sulaeman et al. (2010), with several modifications. Briefly, the experiments were performed using the kit commercialized by BioFilm Control (KITC004) including polystyrene Costar plates with flat bottom (Corning, USA), magnetic beads solution (TON004) and contrast liquid (LIC0001). Two conditions were tested for adhesion assay, microaerobiosis and aerobiosis. Grown cells were recovered from Karmali agar plates and suspended at 108 CFU/mL (OD610 nm = 0.5 ± 0.1) in filtered BHI (provided with the kit). C. jejuni suspensions (200 µL), containing magnetic beads at 1% (vol/vol), were inoculated in Costar plate wells. After 2 h of incubation at 42◦C, the adhesion capability of strains was evaluated by measuring a biofilm formation index (BFI) with the BFC Element 3 software (BioFilm Control, France). Assays were repeated at least three times with three technical replicates.

#### TABLE 1 | Campylobacter jejuni strains experimentally used this study.

\*Cjj: Campylobacter jejuni subsp. jejuni; Cjd: Campylobacter jejuni subsp. doylei; Cj: Campylobacter jejuni.

## Confocal Laser Scanning Microscopy (CLSM)

#### Static Biofilm Formation Assay

Campylobacter jejuni Bf and C. jejuni 81-176 cells were recovered from Karmali agar plates and suspended in BHI at 108 CFU/mL (OD610 nm = 0.5 ± 0.1). Two hundred microliters of bacterial suspension were inoculated in sterile 96 well polystyrene microtiter plates with a micro-clear R bottom 190 ± 5 µm (Greiner Bio One, Germany). Several incubation times (30 min, 1, 2, 4 h) at 42◦C were tested to evaluate the minimum time required for adhesion of the cells at the bottom of the well. Adhesion was performed under microaerobiosis (with bacteria first grown under microaerobiosis) and under aerobiosis (with C. jejuni Bf grown under aerobiosis and C. jejuni 81-176 grown under microaerobiosis). Then, the bacterial suspension in the microtiter plate was carefully replaced with 200 µl of sterile BHI. Plates were then incubated at 42◦C for 24 and 48 h under microaerobic or aerobic conditions. At least 1 h before the biofilm observation, the cells were stained by adding Syto 9 at 0.01 mM final concentration (LIVE/DEAD R Kit, Life Technologies, USA) directly into the wells, following the method of Turonova et al. (2015). Experiments were performed using three biological replicates. For each condition, three technical replicates were performed, and two acquisitions in each of them.

#### Confocal Laser Scanning Microscopy

After staining, image acquisition was performed using a spinning disk confocal microscope (Andor, UK; Olympus, Japan). The entire wells were first inspected to see biofilm formation and its global structure. Two different locations of each well were scanned using a 10X objective lens with the signal recorded in the green channel (excitation 488 nm, emission 500–525 nm). The chosen place for the acquisition was representative of the whole structure and a stack of horizontal planar images with a size of x = 670.8 µm and y = 897.84 µm (e.g., 1040 × 1392 pixels) was scanned with a z-step of 1 µm. Video acquisitions were performed in a selected layer of the same size as described before using a 40X NA 1.4 oil immersion objective lens with an exposure time of 100 ms. Acquisitions were achieved in three distinct positions in the biofilm structure: the bottom, middle and top of the biofilm.

#### Image Processing

confocal laser scanning microscopy (CLSM) images from top to bottom were processed using IMARIS software (v 7.6, Bitplane AG, Switzerland). For visualization of the biofilm, shadow projections and three-dimensional structures were generated. Beside the biofilm appearance, quantitative structural parameters of biofilms were calculated. Biofilm volume and thickness were the selected parameters used to compare the architectural differences of the biofilms formed. The bio-volume corresponds to the total volume of cells in the acquired field (x × y × z = µm3 ) and the thickness is the maximum height reached by the biofilm (µm).

#### Adhesion Assay to Epithelial Intestinal Cells In vitro

Human intestinal cell lines HT29 and HT29-MTX were used to compare adhesion abilities of C. jejuni NCTC 11168, C. jejuni 81- 176 and C. jejuni Bf under microaerobic conditions. In addition, adhesion capabilities of C. jejuni Bf acclimated to ambient air were also assessed. Maintenance of cells and adhesion assays were performed according to Haddad et al. (2010). Briefly, intestinal cells were grown in Dulbecco's minimum essential medium (DMEM) supplemented with 10% fetal bovine serum (FBS), containing 200 mM L-glutamine, 250 µg/mL gentamicin (Sigma-Aldrich, USA) and 2.5 µg/mL amphotericin B (Sigma-Aldrich, USA). The cells were grown routinely in tissue culture flasks at 37◦C in a 5% CO2-humidified atmosphere.

For experimental assays, cultured cells were dissociated from plastic flasks using trypsin-EDTA solution (Invitrogen, USA)

and approximately 105 eukaryotic cells were seeded into each well of 24-well tissues culture tray and incubated for 5 days at 37◦C in humidified atmosphere at 5% of CO2. The cells were washed with DMEM and each well was inoculated with a suspension of approximately 107 CFU of bacteria. To evaluate the number of adhered bacterial cells, the infected monolayers were incubated for 1 h at 37◦C in a humidified 5% CO2 incubator and rinsed five times with phosphate buffered-saline (PBS, Eurobio, France). The cell monolayer was lysed by addition of 0.5 mL of Triton X-100 0.1% (Labo-Si, France) at room temperature for 30 min. C. jejuni cells were enumerated from the lysate on Karmali agar plates after 48 h incubation at 42◦C under microaerobic condition. Experiments were performed using three biological replicates, and for each two technical replicates.

### Genome Sequence Completion and Comparative Genomic Analysis

To complete the draft genome sequence of C. jejuni Bf (Bronnec et al., 2016), PCR amplifications were performed on regions presenting uncertainties and for gap-filling purpose on contig extremities with primers designed in the flanking regions of each gap and PCR products were sequenced (Biofidal, France). As genome comparison showed that C. jejuni Bf was closer to other genomes than that of the reference genome of C. jejuni NCTC 11168 a new mapping was performed on the closest complete genome available (C. jejuni ATCC 32488 SRZ049709). Automatic annotation was performed on the MicroScope platform (MaGe; Vallenet et al., 2006, 2013) and manually checked.

Nucleotide sequence accession number: this whole genome project has been deposited in ENA under the accession no. FCEZ01000001-FCEZ01000095. The version described in this paper is the second version, FCEZ01000001-FCEZ01000095.

Using the tools available on the MicroScope platform, genomic comparisons were conducted between C. jejuni Bf genome and other C. jejuni genomes listed in Supplementary Table S1. A total of 33 complete and 19 draft C. jejuni genomes were used. "PkGDB Synteny Statistics" tool was used to perform similarity analysis between C. jejuni Bf and all C. jejuni genomes available to date on the PkGDB database. "Gene phyloprofile" tool has enabled the genomic comparison by searching specific genes of C. jejuni Bf in comparison with the other genomes, with the following homology constraints: minLrap ≥ 0.8, maxLrap ≥ 0 and identity ≥ 30%.

#### RNA Isolation and Reverse Transcription

After growth AAC or MAC C. jejuni cells were recovered from Karmali plates and suspended in BHI at 108 CFU/mL (OD610 nm = 0.5 ± 0.1). RNA isolation, control and reverse transcription were performed according to Haddad et al. (2012) with some modifications. Briefly, one milliliter of this suspension was centrifuged at 3,300 g for 6 min at 4◦C, and then resuspended in 1 mL of Extract-All (Eurobio, France) and mixed with 0.2 mL of chloroform. After a centrifugation at 12,000 g during 15 min at 4◦C, RNAs from the aqueous phase were precipitated with isopropanol, washed twice in cold 75% ethanol and then solubilized in 50 µL of RNase-free water. Samples were then treated with TurboDNase (Life Technologies, France) to remove potential DNA contamination. The integrity of RNA was verified using 1% agarose gel and quantified using a NanoDrop spectrophotometer (Thermo Scientific, France). Absence of DNA contamination was validated by PCR. RNA was isolated from three biological replicates. Reverse transcription was performed on 100 ng of RNA using the RevertAid H Minus First-Strand cDNA synthesis kit (Euromedex, France) using random hexamer primers according to the manufacturer's instructions.

### Quantitative Real-Time PCR

The quantitative real-time PCR assay was performed using SYBR Green I (Applied Biosystems, USA) and MJ Research PTC-200 Thermal Cycler (GMI, USA). The chosen internal control was rrs (Hyytiäinen et al., 2012) with primers rrs\_F AAGGGCCATGATGACTTGACG and rrs\_R AGCGCAACCCACGTATTTAG. The studied genes were cosR (with primers cosR\_F TTTGAAAGCTGGAGCTGATG and cosR\_R GGTTCCGCCAAGTCTTAGTC) and dnaK (DnaK)\_F AAACGCCAAGCGGTAACTAA and DnaK\_R TTCTTTAGCCGCGTCTTCAT). The operon oorDABC (with primers oorD2\_F TGCGGTTTTAGGACAAATGA and oorD2\_R TTCATCTCTTTTTGCCACCA, oorA2\_F GCGGCAATGAGTGGAGTAAA and oorA2\_R TTGGAAGA CCTGTTGAAGGA, oorB2\_F TGGTAAGTGGAGATGGGG ATA and oorB2\_R GTTGGGCTTGTTTGGGAAT, oorC\_F GTGGTGGCCCTACTAAGGTG and oorC\_R AACCCTTATC TGCAGTCGAAA) was also studied. Finally, a CDS of unknown function (u30002\_F TTCAGAACCTACGAGGATGGA and u30002\_R TTCAATCCTCCAAGCACACA) located upstream from dnaK was also investigated. The PCR mix was prepared as follows: 100 ng to 1 µg of cDNA (for cosR expression or oorDABC, dnaK, and u30002\_F), 1 µM of each primers and 12.5 µL of SYBR Green I Master Mix. The amplification program included an initial denaturing step of 10 min at 95◦C followed by 40 cycles of 15 s at 95◦C and 1 min at 60◦C. A negative control was included in each run. Relative quantification of gene expression was calculated according to the 2−11Ct method (Livak and Schmittgen, 2001). Results were normalized to the gene transcription of the reference strain C. jejuni 81-176 in microaerobic conditions. The experiments were performed in triplicate from three independent cultures. For each experiment, at least three technical replicates were realized.

#### Statistical Analysis

Adhesion results from Biofilm Ring Test were analyzed using Statgraphics Centurion software version 17.1.06 (Statpoint Technologies, USA). An analysis of variance (ANOVA) was assessed to determine the individual effect of each variable (species and atmosphere). Statistical data were completed using the Fisher LSD (least significant difference) technique for multiple comparisons with a significance level at 95%.

Numerical data on biofilm formation obtained from IMARIS were also assessed for an ANOVA. The two variables identified

were the maximum height of biofilm and the biomass volume. The two factors considered were the time of biofilm formation (24 or 48 h) and the combination strain/atmosphere, e.g., C. jejuni 81-176 grown under microaerobiosis (81-176µO2), C. jejuni Bf under microaerobiosis (BfµO2) and C. jejuni Bf under aerobiosis (BfO2). This procedure allows the analysis of variance at several factors for each variable. Significant effects were considered when p-value < 0.05.

Results obtained for the adhesion assay to epithelial intestinal cells in vitro and from RT-qPCR were analyzed using Student's t-test. p-value < 0.05 were considered statistically significant.

#### RESULTS

#### Adhesion Capability and Biofilm Ultrastructure to Abiotic Surfaces Ability to Adhere to Abiotic Surface

Adhesion assays using BioFilm Ring Test R method were conducted under microaerobic and aerobic conditions with an initial bacterial concentration of 5 × 106 CFU/well. According to the biofilm formation index measured with the BFC Element 3 software all strains showed adhesion capacity and could be classified into four groups: strains with strong (0 ≤ BFI < 4), delayed (4 ≤ BFI < 7), or weak adhesion (7 ≤ BFI < 16), and those showing no adhesion capacity (BFI ≥ 16; **Figure 1**).