Introduction

Staphylococcus aureus (S. aureus) is one of the important bacterial pathogens and could cause skin infections, endocarditis, septicemia, meningitis, septic arthritis, toxic shock syndrome, medical device-related infections, gastroenteritis and staphylococcal food poisoning [1, 2]. Staphylococcus aureus produces various virulence factors including enterotoxins, exfoliative toxins, toxic shock syndrome toxin-1, Panton-Valentine leucocidin, and other exoproteins [3]. Staphylococcal food poisoning, one of the most common food-borne diseases in the world, is usually caused by staphylococcal enterotoxins (SEs) [4], which include various types such as SEA, SEB, SEC, SED, SEE, SEG, and SEH [5].

With ever-increasing antimicrobial resistant S. aureus strains (especially methicillin-resistant S. aureus), novel substances, other than antibiotics, to treat and prevent infections caused by this pathogen have been explored. Plant-derived compounds have gained tremendous interest in the search for such alternatives. Extracts from plants have been demonstrated to inhibit different microbial pathogens such as S. aureus, Salmonella, and Listeria [6–8]. The natural compounds responsible for antimicrobial effects include tannins, flavonoids, essential oil, organic acid, and so on [9]. Most of those compounds are derived from plants which have a long history of use in folk medicine or culinary use.

Chlorogenic acid (CA) is a secondary plant metabolite and it is commonly found in plant materials such as apples, coffee beans, and kiwi fruit [10]. CA has antiphlogistic, antioxidant, antimutagenic, and other biological activities [11, 12]. Furthermore, CA has been shown to have antibacterial activity against Trichosporon beigelii, Malassezia furfur, Shigella dysenteriae (S. dysenteriae), Escherichia coli (E. coli), and Salmonella Typhimurium (S. Typhimurium) [13, 14]. Sung et al. [14] have reported that CA exhibits antifungal activities by disrupting the structure of the cell membrane. Lou et al. [13] have found that CA was able to inhibit S. dysenteria and Streptococcus pneumoniae (S. pneumoniae) by provoking irreversible permeability changes in cell membrane, causing cells to lose the ability to maintain membrane potential and cytoplasm macromolecules including nucleotide.

Although CA has been reported to inhibit several pathogens, its effect on S. aureus and its mechanism of action have not been fully explored. Therefore, we investigated the effect of CA on the growth of S. aureus and examined its antistapholococcal mechanism by examining membrane structure under electron microscope and by measuring the intracellular and extracellular ATP concentrations, membrane potential, the intracellular pH (pHin), and the release of cell components.

Materials and methods

Reagents

CA [≥95 %, CAS 327-97-9] was bought from the Sigma Chemical Co. (St. Louis, Mo, USA), and solutions were prepared in 0.9 % (w/v) saline solution containing 1 % (v/v) of DMSO and stored at −20 °C. All other chemicals (including DMSO, HEPES, glycine, citric acid, and glutaraldehyde) were of analytical grade and made by Tianjin Kemiou Chemical Reagent Co., Ltd. (Tianjin, China).

Bacterial strains and culture conditions

Staphylococcus aureus ATCC 25923 was purchased from the American Type Culture Collection (ATCC, Manassas, VA, USA). Seven other S. aureus isolates were taken from our laboratory strain collection and originally isolated from various food sources in China. All isolates were used in minimum inhibitory concentration assay and only S. aureus ATCC 25923 was used in antimicrobial mechanism study. Before each experiment, stock cultures were propagated through two consecutive 12 h growth cycles in LB at 37 °C and washed three times in saline solution. The OD600 of working cultures was 0.5 (containing approximately 107 CFU/ml).

Minimum inhibitory concentration (MIC) determination

The MICs of the CA against S. aureus were determined by the agar dilution method recommended by the European Committee for Antimicrobial Susceptibility Testing [15] with a few modifications. Ampicillin was used as a reference antibiotic. The stock solution of the antibiotic was prepared in sterile water and sterilized with 0.2-μm Acrodisc filter (Gelman, USA). LB agar was aseptically added to sterile glass tubes containing CA or ampicillin. The content (a final volume of 4 ml) of each tube was gently mixed and poured in 24-well plates. The final concentration of CA was 0, 1.25, 2.5, 5, and 10 mg/ml, respectively, and the concentration of ampicillin was 0.1 mg/ml. After hardening, the agar media were spotted with 2 μl (approximately 104 CFU/ml) of the tested bacterium. The spots were left to dry and then plates were incubated at 37 °C for 12 h. The MIC was defined as the lowest concentration (mg/ml) of the extract resulting in no growth of bacteria.

Growth curves

The method described by Qiu et al. [16] was followed. Briefly, S. aureus strain ATCC 25923 was grown to an OD600 value of 0.1 in LB. Then, 150 μl of the culture were placed into wells in 96-well microplates. CA was added to the cultures to obtain final concentrations of 1/32MIC, 1/16MIC, 1/8MIC, 1/4MIC, 1/2MIC and MIC. The S. aureus culture containing 1 % DMSO without CA was the control. Bacteria were further cultured at 37 °C, and cell growth was determined at OD600 at 1 h intervals by a SmartSpec™ Plus Spectrophotometer (Bio-Rad, California, USA).

Cell constituents’ release

The release of cell constituents was measured according to the method described by Rhayour et al. [17]. Cells from the working culture (50 ml) of ATCC 25923 was collected by centrifugation at 4,500 rpm/min for 5 min (Model 5804R, Eppendorf, Hamburg, Germany), washed three times, and resuspended in 0.9 % saline solution at OD600 of 0.5. A 20-ml of cell suspension was incubated at 37 °C for 1 h in the presence of CA at three different concentrations (0, 2MIC, and 4MIC). Then, 10 ml of samples were taken and filtered through a 0.2-μm Acrodisc filter, and then diluted with 0.9 % saline solution. The concentration of the constituents released was determined by a UV spectrophotometer (SmartSpec™ Plus Spectrophotometer, USA) at 260 nm. The absorption of the 0.9 % saline solution containing the same concentration of CA was corrected.

Measurement of intracellular and extracellular ATP concentrations

The method described by Turgis et al. [18] was followed with some modifications. The working culture of ATCC 25923 was centrifuged for 5 min at 4,500 rpm/min, and the supernatant was removed. The cell pellets were washed three times with 0.9 % saline solution and then cells were collected by centrifugation. A cell suspension (OD600 = 0.5) was prepared with 50 ml of 0.9 % saline solution and 4 ml of cell solution was taken into the eppendorf tube for CA treatment. Then, the CA was added to each tube and the final concentrations were 0 (control), MIC, 2MIC, and 4MIC, respectively. Samples were maintained at 37 °C for 30 min and centrifuged for 5 min at 4,500 rpm/min. Then the cells (lower layer) and supernatant (upper layer) were let in ice to prevent ATP loss until measurement. The extracellular and the intracellular ATP concentrations were measured using an ATP assay kit (Nanjing Jiancheng Bioengineering Institute, Nanjing, Jiangsu, China). The ATP concentration of the supernatants, which represents the extracellular concentration, was determined by the kit following instructions by manufacture with a spectrophotometer (SmartSpec™ Plus Spectrophotometer, USA). To determine the intracellular ATP concentrations, cell pellets were washed with 0.9 % saline solution and centrifuged for 5 min at 4,500 rpm/min, and the supernatant was removed. The resulting cell pellets were maintained in 60 μl of 0.9 % saline solution at 100 °C for 10 min to disrupt the microorganisms. The intracellular ATP concentrations were then measured as described for the extracellular ATP concentration.

pHin measurements

pHin was determined according to a modified method of Breeuwer et al. [19]. For loading of cells with fluorescent probe, LB broth (25 ml) was inoculated with 250 μl of working culture (OD600 = 0.5) and incubated at 37 °C for 8 h. Cells were then harvested by centrifugation (4,500 rpm/min, 10 min) and washed twice with 50 mM HEPES buffer (containing 5 mM EDTA, pH 8). The cell pellet was resuspended in 10 ml of HEPES buffer with 1.0 μM fluorescent probe, carboxyfluorescein diacetate succinimidyl ester (cFDA-SE; Molecular Probes, Sigma, USA). Cells were then incubated for 10 min at 37 °C, washed once in 50 mM potassium phosphate buffer with 10 mM MgCl2, pH 7.0, and resuspended in 10 ml buffer. To eliminate nonconjugated cFDA-SE, glucose (10 mM, final concentration) was added and cells were incubated for an additional 30 min at 37 °C. Cells were then washed twice, resuspended in 50 mM phosphate buffer (pH 7), and placed on ice until use.

An aliquot (4 ml) of fluorescently labeled cell suspensions was dispensed in a 15-ml tube with CA at three different concentrations (0, 2MIC and 4MIC). After 30 min, fluorescence intensities were measured with two different excitation wavelengths (440 and 490 nm) by rapidly alternating the monochromator between both wavelengths using the spectrofluorometer (RF-5301, SHIMADZU, Kyoto, Japan). The emission was collected at 525 nm, and the excitation and emission widths were 10 nm for both slits. The pHin of bacteria was determined from the ratio of the fluorescence signal at the pH-sensitive wavelength (490 nm) and the fluorescence signal at the pH-insensitive wavelength (440 nm). During the assay, the system was maintained at 25 °C.

Calibration curves were determined for cFDA-SE-loaded cells with different pH buffers. Controls included the following buffers: glycine (50 mM), citric acid (50 mM), Na2HPO4·2H2O (50 mM), and KCl (50 mM) adjusted to various pH values (4, 5, 6, 7, 8, 9, and 10). The fluorescence intensity was measured at 25 °C after equilibrating the pHin and pHout by addition of valinomycin (1 μM) and nigericin (1 μM). In this assay, a drop in relative fluorescence occurred when the cytoplasmatic pH decreased.

Membrane potential determination

The method described by Sánchez et al. [20] was followed with minor exceptions. Cells were grown in 20 ml of LB broth at 37 °C to an optical density of 0.5 (approximately 107 CFU/ml) at 600 nm. Then, 1 μM of membrane potential sensitive fluorescent probe bis-(1,3-dibutylbarbituric acid) trimethine oxonol [DiBAC4(3); Molecular Probes, Sigma, USA] was added for 5 min, followed by the addition of CA (0, 2MIC, and 4MIC). After 5 min, fluorescence was measured at the excitation and emission wavelengths of 492 nm and 515 nm, respectively, using the spectrofluorometer (RF-5301, Japan). The excitation and emission slit widths were 3 and 5 nm, respectively. Background fluorescence resulting from the medium was corrected.

Field emission scanning electron microscope (FESEM) analysis

FESEM assays were carried out as previously reported [21]. Cells (OD600 = 0.5) were treated with CA at 0, 2MIC, and 4MIC, respectively. The samples were incubated at 37 °C. After incubation for 2 or 4 h, cells were harvested by centrifugation for 10 min at 4,500 rpm/min and washed twice with 0.9 % saline solution, and then were resuspended in water containing 2.5 % glutaraldehyde and kept at −4 °C for 10 h to fix the cells. After centrifugation, the cells were further dehydrated in water–alcohol solutions at various alcohol concentrations (30, 50, 70, 80, 90, and 100 %) for 10 min. Finally, the samples were fixed on CLSM support, and then sputter coated with gold under vacuum, followed by microscopic examinations using a field emission scanning electron microscope (S-4800, Hitachi, Tokyo, Japan).

Statistical analysis

All experiments were performed in triplicate. Statistical analyses were performed using the SPSS software (version 19.0; SPSS, Inc., Chicago, IL). The data were presented as the mean values ± SD (n = 6). Results were analyzed with an analysis of variance test, and an independent Student’s t test was used for the analysis. Differences between means were considered significant at P ≤ 0.05.

Results and discussion

Growth inhibitory effect of CA on S. aureus

The MICs of CA against eight S. aureus strains were determined and ranged from 2.5 to 5.0 mg/ml. The MIC of CA against S. aureus ATCC 25923 was found to be 2.5 mg/ml (Table 1). At the concentrations of 1/16, 1/8, 1/4, and 1/2MIC, CA had no obvious effects on the growth of S. aureus. However, the growth of S. aureus was attenuated remarkably in the presence of CA treatment at concentrations of MIC or 2MIC. The OD600 values of treated culture with CA at these two concentrations were 40–60 % of the untreated culture after 9 h (Fig. 1).

Table 1 Minimum inhibitory concentrations of chlorogenic acid against different strains of S. aureus
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Fig. 1
figure 1

Growth curves for S. aureus ATCC 25923 cultured in LB with various concentrations of chlorogenic acid. Each value represents the average of three reproducible experiments

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Cell constituents’ release

The cell constituents’ release was determined by the measurement of the absorbance at 260 nm of the filtrates of S. aureus cultures. After CA was added to S. aureus, a significant (P ≤ 0.05) increase in the cell constituents’ release was observed when S. aureus were treated with CA at 2MIC. The absorbance was 0.18 ± 0.05 for CA (2MIC)-treated cells, while it was 0.04 ± 0.01 for the control. When the concentration of CA increased from 2MIC to 4MIC, the release of cell constituents increased gradually with the increase in the CA’s concentration (Fig. 2).

Fig. 2
figure 2

Effect of chlorogenic acid on cell constituents’ release of S. aureus ATCC 25923. Values represent the average of three reproducible experiments. Bars represent the standard deviation (n = 6). *P < 0.05, **P < 0.01

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A loss of 260-nm absorbing materials has been observed after CA treatments. Compounds, such as oil, oregano, carvacrol, nobiletin, and tangeretin, induced a loss of cytoplasmic constituents inside of bacteria including K+, nucleic acids, protein, and ATP [7, 18, 22, 23]. Ultee et al. [24] demonstrated that 0.25 and 1 mM carvacrol induced release of potassium ions in Bacillus cereus (B. cereus). Lv et al. [21] found that essential oil, derived from ten types of plants including Pogostemon cablin, Salvia sclarea, and Rosmarinus officinalis, damaged the membrane structure of Pseudomonas aeruginosa (P. aeruginosa) with the significant increase in the cell constituents’ release. Yao et al. [22] observed that nobiletin and tangeretin destroyed the permeability of the membrane of P. aeruginosa and S. aureus, with release of alanine transaminase and aspartate transaminase. The release of those constituents indicated that the membrane structure of microorganism has been damaged by natural compounds.

Intracellular and extracellular ATP concentrations

The intracellular and extracellular ATP concentrations in the untreated S. aureus cells were 75.25 ± 6.00 and 9.50 ± 2.10 pM, respectively (Fig. 3). Compared with control, S. aureus treated with CA from MIC to 4MIC showed a significant (P ≤ 0.05) reduction in intracellular ATP concentration, while no significant (P > 0.05) difference was observed in the extracellular ATP concentration. When the concentration of CA increased from MIC, 2MIC to 4MIC, intracellular ATP concentrations were 4.50 ± 4.40, 7.35 ± 3.7, and 2.25 ± 1.8 pM, respectively, and extracellular ATP concentrations were 6.50 ± 2.10, 9.50 ± 0.7, and 8.50 ± 0.7 pM, respectively. However, no significant (P > 0.05) difference was observed in the intracellular and extracellular ATP concentration by increasing the concentration of CA from MIC to 4MIC.

Fig. 3
figure 3

Effect of chlorogenic acid on intracellular and extracellular ATP concentrations of S. aureus ATCC 25923. Values represent the means of three reproducible experiments. Bars represent the standard deviation (n = 6). *P < 0.05, **P < 0.01

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We found that exposure to CA led to a decrease in the intracellular ATP concentration. No proportional increase in the extracellular ATP was observed. It is possible that CA does not enhance the permeability of the membrane for ATP, but it could interact with the membranes of S. aureus and change its permeability for cations like H+ and K+. Depletion of the internal ATP pool may result from a reduced rate of ATP synthesis or increased ATP hydrolysis. Our results were consistent with Ultee et al. [24] who reported that the level of intracellular ATP decreased, while there was no proportional increase outside the cell after B. cereus cell was exposed to carvacrol. In contrast, Lee et al. [25] observed that linolenic acid with or without monoglyceride (glycerol laurate or glycerol myristate) changed both the intracellular and extracellular ATP concentrations of B. cereus and S. aureus. Turgis et al. [18] reported that mustard essential oil induced a decrease in the intracellular ATP concentration and increased the extracellular ATP concentration. The differences in the permeability of the cell membrane caused by natural compounds may be due to the various degrees of damages of cell membranes which are dependent on the membrane structure of pathogens and antimicrobial effects of different phytochemicals.

pHin

A clear change in cytoplasmic pH was observed after addition of CA (Fig. 4). The pHin of untreated S. aureus was 9.22 ± 0.77. The addition of CA at 2MIC caused a significant (P ≤ 0.05) decrease in pHin of S. aureus from 9.22 ± 0.77 to 5.71 ± 1.07. And the addition of CA at 4MIC significantly (P ≤ 0.05) lowered the pHin from 9.22 ± 0.77 to 4.50 ± 0.72. However, there was no significant (P > 0.05) difference in pHin by increasing the concentration of CA from 2MIC to 4MIC.

Fig. 4
figure 4

Effect of chlorogenic acid on the intracellular pH (pHin) of S. aureus ATCC 25923. Values represent the means of three reproducible experiments. Bars represent the standard deviation (n = 6). *P < 0.05, **P < 0.01

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The pHin is critical for the control of many cellular processes, such as DNA transcription, protein synthesis, and enzyme activity and also controlled by the membrane [26]. Changes in pHin are an indicator of membrane integrity. Turgis et al. [18] demonstrated that mustard essential oil at concentration of the MIC lowered the pHin from 6.23 to 5.20 in E. coli O157:H7 and from 6.59 to 5.44 in S. typhi. Sanchez et al. [20] showed that all the plant extracts induced changes in the pHin of Vibrio cholerae (V. cholerae). The results indicated that plants compounds generally damaged the membrane integrity and caused a decrease in pHin.

Membrane potential

Staphylococcus aureus cells treated with CA displayed cell membrane hyperpolarization, as evidenced by a decrease in fluorescence (negative values) (Fig. 5). After the addition of CA at 2MIC or 4MIC, the membrane potential was decreased, and a higher rate of reduction was observed by increasing concentrations from 2MIC to 4MIC.

Fig. 5
figure 5

Effect of chlorogenic acid on the membrane potentials of S. aureus ATCC 25923. Values represent the means of three reproducible experiments. Bars represent the standard deviation (n = 6)

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A reduction in fluorescence and a hyperpolarization of cellular membranes have been observed after CA treatment. The fluorescence emitted by DiBAC4(3), a fluorescent membrane potential dye, is enhanced when the dye crosses the cell membrane as a result of membrane depolarization (less negative charge inside the cell). Lou et al. [13] found that CA caused depolarization of cell membranes of S. dysenteria and S. pneumoniae. Although a depolarization effect was expected, hyperpolarization has also been reported as an important type of membrane damage [27]. Consistent with our results, Sanchez et al. [20] reported that extracts from edible and medicinal plants were able to disrupt the cell membranes of V. cholerae cells, causing hyperpolarization of cell membrane. There are two reasons to explain the phenomenons: (1) a pH change (from acidic to neutral); and (2) by increasing movement of ions, specifically K+, which diffuses out of the cell membrane through K+ channels and affects cellular homeostasis [28].

FESEM observation

Cells of S. aureus had a typical structure of gram-positive microorganism, showing an irregular and striated membrane (Fig. 6). Cells treated with CA showed disrupted membranes and deformed cell walls. Cells treated with higher concentrations of CA or longer time exhibited more damage to cell membrane than those untreated cells.

Fig. 6
figure 6

Scanning electron micrographs of S. aureus ATCC 25923 untreated for 2 h (a) and 4 h (b), treated with chlorogenic acid at 2MIC for 2 h (c) and 4 h (d), and treated with chlorogenic acid at 4MIC for 2 h (e) and 4 h (f)

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Microstructural observations showed that the cell membrane structures of cells were severely affected and damaged by CA. The findings are in good agreement with other studies. Observed by scanning electron microscope, Rhayour et al. [17] demonstrated that cell walls and membranes of E. coli and B. subtilis treated by oregano and clove essential oils were significantly damaged. Turgis et al. [18] reported that a variety of morphological changes in E. coli O157:H7 and S. typhi caused by the mustard EO had been observed by SEM.

Conclusion

In conclusion, CA effectively inhibited the growth of S. aureus partially by affecting the cell membrane, which was evidenced by the damaged membrane structure, the release of cytoplasmic constituents, and changes in the pHin and membrane potential. The findings reveal the potential of CA as an alternative therapeutic agent against S. aureus or as a food preservative. Prior to the application of CA in either area, issues of safety and toxicity of CA need to be evaluated in future studies.