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Low pH-induced membrane fatty acid alterations in oral bacteria
FEMS Microbiology Letters 238 (2004) 291–295 www.fems-microbiology.org
Low pH-induced membrane fatty acid alterations in oral bacteria Elizabeth M. Fozo a, Jessica K. Kajfasz a, Robert G. Quivey Jr.
a,b,*
a
b
Department of Microbiology and Immunology, University of Rochester, Rochester, NY, 14642, USA Center for Oral Biology in the Aab Institute for Biomedical Research, University of Rochester, P.O. Box 611, 601 Elmwood Avenue, Rochester, NY 14642, USA Received 25 May 2004; received in revised form 15 July 2004; accepted 20 July 2004 First published online 29 July 2004
Abstract Four oral bacterial strains, of which two are considered aciduric and two are considered acid-sensitive, were grown under glucose-limiting conditions in chemostats to determine whether their membrane fatty acid profiles were altered in response to environmental acidification. Streptococcus gordonii DL1, as well as the aciduric strains S. salivarius 57.I, and Lactobacillus casei 4646 increased the levels of mono-unsaturated membrane fatty acids. The non-aciduric strain S. sanguis 10904 did not alter its membrane composition in response to pH values examined here. Thus, in response to low pH, aciduric oral bacteria alter their membrane composition to contain increased levels of long-chained, mono-unsaturated fatty acids. This suggests that membrane fatty acid adaptation is a common mechanism utilized by bacteria to withstand environmental stress. Ó 2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. Keywords: Oral bacteria; Acid adaptation; Membrane fatty acids
1. Introduction During growth in dental plaque, oral bacteria experience cycles of acidification of their environment, followed by neutralization periods [1]. Oral streptococci employ numerous mechanisms to tolerate these periods of low pH, including ammonia-generating activities, the proton-pumping activity of F-ATPase, as well the upregulation of DNA and protein repair systems [2]. Recently, it was shown that membrane fatty acid adaptation was necessary for Streptococcus mutans to survive low pH environments [3]. In response to acidification, S. mutans increases the proportion of the long-chained, mono-unsaturated fatty acids (C18:1 and C20:1) in its membrane with a concomitant decrease in short* Corresponding author. Tel.: +1 585 275 0382; fax: +1 585 276 0190. E-mail address: [email protected] (R.G. Quivey Jr.).
chained, saturated fatty acids (C14:0 and C16:0) as a result of acidification [3]. Although the membrane fatty acid contents of other oral bacteria have been examined, a detailed examination of membrane fatty acid alterations in response to environmental acidification has not been performed [4–10]. Thus, in order to determine whether membrane fatty acid adaptation is a unique survival mechanism for S. mutans in the oral cavity or a more global, bacterial response, we examined whether this phenomenon occurs in other oral bacteria. The oral bacteria S. sanguis 10904, S. gordonii DL1, S. salivarius 57.I, and Lactobacillus casei 4646 were grown under steady-state conditions in a chemostat at neutral and acidic pH values. The membrane fatty acid content was then determined by GC-FAME by Avanti Polar Lipids, Inc. (Alabaster, AL). There were alterations of varying degrees observed in the organisms S. gordonii, S. salivarius, and L. casei during growth at low pH, whereas the non-aciduric S. sanguis did not alter its membrane content substantially. These results suggest
0378-1097/$22.00 Ó 2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. doi:10.1016/j.femsle.2004.07.047
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that membrane fatty acid adaptation is a common mechanism utilized by bacteria to survive acidic environments.
2. Materials and methods 2.1. Strains and growth conditions S. salivarius 57.I [11–13], S. gordonii DL1 [14] (a generous gift from R.A. Burne; University of Florida), and L. casei ATCC4646 [15] were maintained on brain heart infusion (BHI; Difco) medium. S. sanguis NTCC10904 [16–20] was maintained on Todd Hewitt (TH; Difco) medium. All strains were grown on solid medium at 37 °C in an atmosphere containing 5% (vol/vol) CO2, 95% air. Continuous cultures were grown in a Sixfors multiple fermentor system (ATR; Laurel, MD). All strains exept S. sanguis, were grown in TY medium (3% tryptone, 0.1% yeast extract, 0.5% KOH, 1 mM H3PO4) and were maintained at steady-state by glucose limitation as described previously [3]. S. sanguis was grown in TH broth. Culture pH values were maintained by automated addition of 2 N KOH. S. salivarius and L. casei were grown at steady-state at pH 7 and 5; S. sanguis and S. gordonii were grown at pH 7 and 6, as they were unable to grow at pH 5 (data not shown). All streptococcal strains were grown at a dilution rate of 0.24 h 1; L. casei at a dilution rate of 0.098 h 1. Aliqouts (100 ml) of each strain, grown at each pH condition, were harvested via centrifugation and stored at 80 °C for fatty acid determination. Batch cultures (100 ml) were grown overnight in TY with 1% glucose, or in the case of S. sanguis, in TH. Samples of both TH and TY broth were analyzed to determine whether they contained fatty acids that may interfere with interpretation of the data. There were no fatty acids present at the level of detection used in this analysis (data not shown) [21]. Two Escherichia coli strains (a generous gift from J.E. Cronan Jr.; University of Illinois) were used as positive and negative controls for the presence of cylcopropane fatty acids as described previously [21]. Both strains were grown in Luria–Bertani broth until late-log phase at 37 °C, shaking at 250 rpm. Strain ZK126 served as the positive control for the production of cyclopropane fatty acids; strain YYC1272 served as a negative control for cyclopropane fatty acid production [22]. 2.2. Membrane fatty acid determinations Total lipids (approximately 5 mg obtained per sample) were extracted using the method of Bligh [23]. Fatty acid esters were prepared and gas chromatography of fatty acid methyl esters were performed by Avanti Polar Lipids, Inc. (Alabaster, AL), as previously described [3,21].
2.3. Statistics Each strain was grown independently at least twice at each condition. Data are shown as the averages of the percentage of total membrane fatty acids ± standard deviations. StudentÕs t-test was performed as indicated to determine statistical significance. In addition, StudentÕs t-test was used to make pair-wise comparisons in the streptococci strains between the (C14:0 + C16:0)/ (C18:1 + C20:1) ratios found in the cultures. We chose this ratio for comparisons, based on the observations made here and in our previous work with S. mutans UA159. These fatty acids dominate the membrane composition, and in response to environmental acidification, their proportions change significantly [3,21]. Moreover, we have recently shown that there is a direct correlation between this ratio and acid survival for S. mutans [24]. In the case of L. casei, pair-wise comparisons were made between the (C16:0 + C17:0cpa)/(C18:1 + C19:0cpa), as these fatty acids dominated the membrane composition under each growth condition examined.
3. Results and discussion Alterations in the membrane fatty acid or phospholipid composition are common adaptation mechanisms for many bacteria in response to environmental stress. The cariogenic organism S. mutans alters its membrane fatty acid profile in response to acidification of its environment, specifically by increasing the proportions of monounsaturated membrane fatty acids when grown in low pH environments [3,21]. If the ability to increase or produce mono-unsaturated fatty acids is prevented, the organism is rendered acid sensitive [3,24]. Since membrane fatty acid adaptation is a protective measure utilized by S. mutans, it is possible that other oral bacteria utilize this mechanism to protect against acidification of the environment. We grew four oral bacteria, both aciduric organisms and those considered acid-sensitive, in glucose-limited chemostats under controlled pH conditions. Cells were harvested and membrane fatty acids were determined by GC-FAME by Avanti Polar Lipids, Inc. Although S. sanguis has demonstrated a weak ability to adapt to acidification [17,20,25,26], it is not considered aciduric and is not as capable as S. mutans in survival of low pH environments [15,18,19,27,28]. Specifically, studies have shown that S. sanguis 10904 possesses an F-ATPase, a necessary component for low pH survival of oral streptococci, which is not as active at pH values as low as those reported for the S. mutans enzyme [13,15,18–20]. Since it is not considered aciduric and its F-ATPase does not function at pH values as low as its aciduric counterpart in S. mutans, we investigated whether the membrane fatty acid profile of S. sanguis would alter as a response to acidification.
E.M. Fozo et al. / FEMS Microbiology Letters 238 (2004) 291–295
As shown, the membrane fatty acid content primarily consisted of C16:0, C16:1, and C18:1 (Table 1), and did not change significantly in response to pH. However, when grown in batch cultures, there were significant increases in the proportion of C18:1, similar to those reported for S. mutans [21,24]. In addition, the batch culture pH of S. sanguis was approximately 5.5, lower than the pH for chemostat growth. It may be possible that S. sanguis is capable of membrane fatty acid alterations when the environmental pH is lowered to a critical value. S. gordonii, a member of the sanguis group associated with bacterial endocarditis undergoes phenotypic acid adaptation, but is typically not considered aciduric or cariogenic [14,20,26,29]. As shown in Table 1, the organism increased the proportion of the membrane fatty acid C18:1 when the culture pH was lowered from a value of 7 to 6. Although we were unable to grow the organism in steady-state at pH 5, the pH of the batch cultures was approximately 4.7 after 16 h of growth (data not shown). The batch cultures showed even higher levels of C18:1 than those observed in the pH 6 steady-state cultures. Again, it is possible that these elevated levels are due to further acid adaptation of the batch culture than the chemostat culture. It has been shown that particular strains of oral streptococci can undergo phenotypic acid adaptation when grown with excess glucose, similar to the batch conditions used here [17]. Additionally, although both S. gordonii and S. sanguis are not considered aciduric, it is interesting to note that S. gordonii altered its membrane fatty acid profile in response to growth at pH 6 in steady-
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state, unlike S. sanguis. Although the strains were grown in different media, neither media contained detectable levels of fatty acids (data not shown). Thus, there appears to be a difference in the regulation of the fatty acid composition between these two strains. S. salivarius is considered aciduric and can rapidly adapt to acidification of its environment [13,26]. The membrane fatty acid composition of pH 7-grown S. salivarius consisted primarily of C14:0 and C16:0 fatty acids similar to S. mutans [21] (72% of the total membrane fatty acid composition in S. salivarius, 62% of the total in S. mutans). When grown at pH 5, the levels of C14:0 and C16:0 decreased dramatically to comprise only 38.6% of the total composition (Table 1); there was a simultaneous increase in the levels of C18:1 and C20:1, similar to what was reported in S. mutans [21]. The batch-grown cells had membrane fatty acid profiles similar to the pH 5-grown chemostat cells and to reports published previously [4,6–9]. Thus S. salivarius, like S. mutans, increased the proportion of mono-unsaturated membrane fatty acids in response to growth under low pH [21]. Previously, we have demonstrated that if increases in the mono-unsaturated fatty acid composition in S. mutans are blocked, the organism is rendered highly acid-sensitive [3]. It is highly possible that these increases in mono-unsaturated membrane fatty acids are necessary for S. salivarius to survive the acidic environment of dental plaque. L. casei is considered to be extremely aciduric [13,15,19,26,30], and previous reports have demonstrated that Lactobacillus species produce membranes
Table 1 Membrane fatty acid composition of S. sanguis 1 0904, S. gordonii DL 1, S. salivarius 57.1 grown at steady-state in a chemostata Fatty acid
% of total membrane fatty acid composition S. sanguis 10904
C14:0 C14:1 C16:0 C16:1 C18:0 C18:1 C20:0 C20:1 Others C14:0 + C16:0 C18:1 + C20:1 Ratioc
S. gordonii DL 1 b
S salivarius 57.1 b
pH 7
pH 6
Batch (5.5)
pH 7
pH 6
Batch (4.7)
pH 7
pH 5
Batch (4.3)b
6.1 ± 10.1 1.4 ± 0.0 18.1 ± 0.8 27.7 ± 0.6 2.2 ± 0.1 33.6 ± 1.8 ND ND 11.0 ± 2.0 24.2 ± 0.7 33.6 ± 1.8 0.7 ± 0.0
4.8 ± 0.0* 1.6 ± 0.3 14.8 ± 1.1 28.9 ± 1.7 2.3 ± 0.6 33.5 ± 0.1 ND ND 14.1 ± 0.1 19.6 ± 1.1* 33.5 ± 1.1 0.6 ± 0.0*
3.9 ± 0.1* 0.7 ± 0.4 17.6 ± 0.6 23.1 ± 2.1 4.2 ± 0.2* 43.6 ± 2.2* ND 0.1 ± 0.1 7.1 ± 3.0 21.5 ± 0.8* 43.7 ± 2.3* 0.5 ± 0.0*
21.3 ± 0.1 1.4 ± 0.0 44.3 ± 0.1 12.1 ± 0.1 3.7 ± 0.1 11.3 ± 0.1 ND ND 6.1 ± 0.2 65.5 ± 0.1 11.3 ± 0.4 5.8 ± 0.1
15.8 ± 1.5* 1.1 ± 0.0 41.5 ± 0.8* 13.2 ± 0.6 4.9 ± 0.4 19.4 ± 1.6* ND ND 4.3 ± 0.4 57.2 ± 2.3* 19.4 ± 1.6* 3.0 ± 0. 4*
4.6 ± 0.1* 0.7 ± 0.1 29.9 ± 1.4* 6.2 ± 0.6* 16.1 ± 0.9* 40.1 ± 1.8* 0.2 ± 0.1 0.2 ± 0.1 2.3 ± 0.2 34.5 ± 1.5* 40.3 ± 1.9* 0.9 ± 0.1*
20.2 ± 4.2 ND 52.3 ± 1.2 8.7 ± 0.3 5.7 ± 0.9 8.8 ± 2.2 0.1 ± 0.1 1.5 ± 0.8 2.9 ± 0.4 72.2 ± 3.0 10.2 ± 3.0 7.5 ± 2.5
6.3 ± 3.8 ND 32.3 ± 2.0* 4.3 ± 2.0 12.6 ± 5.6 23.1 ± 0.9* 3.3 ± 0.2* 16.0 ± 2.1* 2.3 ± 0.6 38.6 ± 5.8* 39.1 ± 3.0* 1.0 ± 0.2
5.9 ± 0.3* ND 37.6 ± 0.1* 6.6 ± 1.2 7.6 ± 0.4 28.2 ± 1.6* 0.6 ± 0.0* 12.4 ± 0.8* 1.3 ± 0.4 43. 5 ± 0.4* 40.6 ± 2.3* 1.1 ± 0.1
ND, not detected. Average % fatty acid ± standard deviation. n = 2. a Each strain was grown in a chemostat at least two times at a dilution rate of 0.24 h 1. Aliquots were collected after 10 generations at each pH value. b Batch cultures were grown overnight at least two times in an environment enriched with 5% CO2. pH of the batch cultures after 16 h of growth is indicated. c Ratio is (C14:0 + C16:0)/(C18:1 + C20:1). * Statistically significant difference between that condition and pH 7 value, p < 0.05.
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Table 2 Membrane fatty acid composition of L. casei 4646a Fatty acid
C14:0 C14:1 C16:0 C16:1 C17:0 cpa C18:0 C18:1 C19:0 cpa C20:0 C20:1 Others C16:0 ± C17:0 cpa C18:1 ± Cl9;0 cpa Ratioc
% of total membrane fatty acid composition pH 7
pH 5
Batch(3.8)b
3.0 ± 0.1 0.3 ± 0.1 42.7 ± 0.1 4.8 ± 0.4 21.4 ± 0.4 0.9 ± 0.0 11.8 ± 1.3 10.2 ± 0.8 0.0 ± 0.0 0.0 ± 0.0 5.0 ± 0.6 64.1 ± 0.6 22.0 ± 0.5 2.9 ± 0.1
3.7 ± 1.1 0.3 ± 0.1 30.4 ± 2.6* 7.9 ± 2.3 2.7 ± 0.3* 2.6 ± 0.8 25.3 ± 0.1* 25.1 ± 0.6* 0.0 ± 0.0 0.0 ± 0.0 2.0 ± 0.4 33.1 ± 2.3* 50.3 ± 0.6* 0.7 ± 0.0*
4.4 ± 0.6 0.6 ± 0.8 21.2 ± 8.5 16.2 ± 4.6 0.0 ± 0.0* 1.7 ± 0.2 25.9 ± 2.9* 18.6 ± 3.4 0.0 ± 0.0 0.0 ± 0.0 11.5 ± 4.2 21.2 ± 8.5* 44.5 ± 0.5* 0.5 ± 0.2*
ND, Not Detected Average % fatty acid ± standard deviation, n = 2. a L casei was grown in a chemostat two times at a dilution rate of 0.1 h 1. Aliquots were collected after 10 generations at each pH value. b Batch cultures were grown two times in an environment enriched with 5% CO7. Indicated is the pH of the cultures after 30 h of growth. c Ratio is (C16:0 + C17:0cpa)/ (C18:1 + C19:0cpa). * Statistically significant between that condition and pH 7 value, p < 0.05.
that contain cyclopropane fatty acids [5,10]. We examined the membrane fatty acid content of L. casei 4646 after growth at neutral and acidic pH values to determine whether the aciduric bacterium altered its membrane fatty acid content. As shown in Table 2, L. casei does alter its fatty acid content in response to acidification. L. casei, similar to S. mutans, S. gordonii, and S. salivarius increased the proportion of C18:1 in response to acidification, again suggesting that increased levels of longchained, monounsaturated fatty acids are necessary for low pH survival and is a mechanism commonly used by bacteria. In addition, L. casei increased the proportion of cyclopropane fatty acid C19:0 with a concomitant decrease in the levels of C16:0 and cyclopropane fatty acid C17:0. While the relative total amounts of cyclopropane fatty acids did not change with respect to growth pH, the proportion of long versus shorter chained cyclopropane fatty acids did alter. It appears that increased fatty acid length is another important membrane alteration to increase survival in acidic environments. The necessity for cyclopropane fatty acids in low pH survival of E. coli has been demonstrated, and it is likely that they are significant for the survival of L. casei as well [22,31]. In addition, cyclopropane fatty acids have also been associated with disease persistence in Mycobacterium tuberculosis and with the infectious dose of E. coli O157:H7 [32,33]. L. casei is considered to be more aciduric than S. mutans and the other oral streptococci, so the presence of cyclopropane fatty acids may increase the acid tolerance of the lactobacilli, although this remains to be determined. We have demonstrated that the oral bacteria S. gordonii. S. salivarius, and L. casei undergo membrane fatty acid alterations as a result of environmental acidification.
This may be an additional mechanism utilized by the bacteria to increase their survival during the repeated cycles of acidification in dental plaque. Recent work with S. mutans suggest that monounsaturated membrane fatty acids are necessary for maintenance of DpH across the membrane, and it is likely that this function is conserved among the species examined here [24]. It seems likely that other bacteria that must survive pH fluctuations undergo similar membrane changes. Membrane fatty acid alterations may represent a generalized response utilized by many bacteria to survive environmental stress. New therapeutics designed to target the mechanisms responsible for membrane fatty acid adaptation could increase the current arsenal employed to combat disease.
Acknowledgements The authors wish to thank Robert Marquis for helpful discussion throughout. We also thank Roberta Faustoferri for technical expertise. This work was supported by grants from the NIH, National Institute for Dental and Craniofacial Research DE-11549 and DE-06127. E.M.F and J.K.K. were supported by the Rochester Training Program in Oral Infectious Diseases, T32-DE07165.
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Low pH-induced membrane fatty acid alterations in oral bacteria Elizabeth M. Fozo a, Jessica K. Kajfasz a, Robert G. Quivey Jr.
a,b,*
a
b
Department of Microbiology and Immunology, University of Rochester, Rochester, NY, 14642, USA Center for Oral Biology in the Aab Institute for Biomedical Research, University of Rochester, P.O. Box 611, 601 Elmwood Avenue, Rochester, NY 14642, USA Received 25 May 2004; received in revised form 15 July 2004; accepted 20 July 2004 First published online 29 July 2004
Abstract Four oral bacterial strains, of which two are considered aciduric and two are considered acid-sensitive, were grown under glucose-limiting conditions in chemostats to determine whether their membrane fatty acid profiles were altered in response to environmental acidification. Streptococcus gordonii DL1, as well as the aciduric strains S. salivarius 57.I, and Lactobacillus casei 4646 increased the levels of mono-unsaturated membrane fatty acids. The non-aciduric strain S. sanguis 10904 did not alter its membrane composition in response to pH values examined here. Thus, in response to low pH, aciduric oral bacteria alter their membrane composition to contain increased levels of long-chained, mono-unsaturated fatty acids. This suggests that membrane fatty acid adaptation is a common mechanism utilized by bacteria to withstand environmental stress. Ó 2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. Keywords: Oral bacteria; Acid adaptation; Membrane fatty acids
1. Introduction During growth in dental plaque, oral bacteria experience cycles of acidification of their environment, followed by neutralization periods [1]. Oral streptococci employ numerous mechanisms to tolerate these periods of low pH, including ammonia-generating activities, the proton-pumping activity of F-ATPase, as well the upregulation of DNA and protein repair systems [2]. Recently, it was shown that membrane fatty acid adaptation was necessary for Streptococcus mutans to survive low pH environments [3]. In response to acidification, S. mutans increases the proportion of the long-chained, mono-unsaturated fatty acids (C18:1 and C20:1) in its membrane with a concomitant decrease in short* Corresponding author. Tel.: +1 585 275 0382; fax: +1 585 276 0190. E-mail address: [email protected] (R.G. Quivey Jr.).
chained, saturated fatty acids (C14:0 and C16:0) as a result of acidification [3]. Although the membrane fatty acid contents of other oral bacteria have been examined, a detailed examination of membrane fatty acid alterations in response to environmental acidification has not been performed [4–10]. Thus, in order to determine whether membrane fatty acid adaptation is a unique survival mechanism for S. mutans in the oral cavity or a more global, bacterial response, we examined whether this phenomenon occurs in other oral bacteria. The oral bacteria S. sanguis 10904, S. gordonii DL1, S. salivarius 57.I, and Lactobacillus casei 4646 were grown under steady-state conditions in a chemostat at neutral and acidic pH values. The membrane fatty acid content was then determined by GC-FAME by Avanti Polar Lipids, Inc. (Alabaster, AL). There were alterations of varying degrees observed in the organisms S. gordonii, S. salivarius, and L. casei during growth at low pH, whereas the non-aciduric S. sanguis did not alter its membrane content substantially. These results suggest
0378-1097/$22.00 Ó 2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved. doi:10.1016/j.femsle.2004.07.047
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that membrane fatty acid adaptation is a common mechanism utilized by bacteria to survive acidic environments.
2. Materials and methods 2.1. Strains and growth conditions S. salivarius 57.I [11–13], S. gordonii DL1 [14] (a generous gift from R.A. Burne; University of Florida), and L. casei ATCC4646 [15] were maintained on brain heart infusion (BHI; Difco) medium. S. sanguis NTCC10904 [16–20] was maintained on Todd Hewitt (TH; Difco) medium. All strains were grown on solid medium at 37 °C in an atmosphere containing 5% (vol/vol) CO2, 95% air. Continuous cultures were grown in a Sixfors multiple fermentor system (ATR; Laurel, MD). All strains exept S. sanguis, were grown in TY medium (3% tryptone, 0.1% yeast extract, 0.5% KOH, 1 mM H3PO4) and were maintained at steady-state by glucose limitation as described previously [3]. S. sanguis was grown in TH broth. Culture pH values were maintained by automated addition of 2 N KOH. S. salivarius and L. casei were grown at steady-state at pH 7 and 5; S. sanguis and S. gordonii were grown at pH 7 and 6, as they were unable to grow at pH 5 (data not shown). All streptococcal strains were grown at a dilution rate of 0.24 h 1; L. casei at a dilution rate of 0.098 h 1. Aliqouts (100 ml) of each strain, grown at each pH condition, were harvested via centrifugation and stored at 80 °C for fatty acid determination. Batch cultures (100 ml) were grown overnight in TY with 1% glucose, or in the case of S. sanguis, in TH. Samples of both TH and TY broth were analyzed to determine whether they contained fatty acids that may interfere with interpretation of the data. There were no fatty acids present at the level of detection used in this analysis (data not shown) [21]. Two Escherichia coli strains (a generous gift from J.E. Cronan Jr.; University of Illinois) were used as positive and negative controls for the presence of cylcopropane fatty acids as described previously [21]. Both strains were grown in Luria–Bertani broth until late-log phase at 37 °C, shaking at 250 rpm. Strain ZK126 served as the positive control for the production of cyclopropane fatty acids; strain YYC1272 served as a negative control for cyclopropane fatty acid production [22]. 2.2. Membrane fatty acid determinations Total lipids (approximately 5 mg obtained per sample) were extracted using the method of Bligh [23]. Fatty acid esters were prepared and gas chromatography of fatty acid methyl esters were performed by Avanti Polar Lipids, Inc. (Alabaster, AL), as previously described [3,21].
2.3. Statistics Each strain was grown independently at least twice at each condition. Data are shown as the averages of the percentage of total membrane fatty acids ± standard deviations. StudentÕs t-test was performed as indicated to determine statistical significance. In addition, StudentÕs t-test was used to make pair-wise comparisons in the streptococci strains between the (C14:0 + C16:0)/ (C18:1 + C20:1) ratios found in the cultures. We chose this ratio for comparisons, based on the observations made here and in our previous work with S. mutans UA159. These fatty acids dominate the membrane composition, and in response to environmental acidification, their proportions change significantly [3,21]. Moreover, we have recently shown that there is a direct correlation between this ratio and acid survival for S. mutans [24]. In the case of L. casei, pair-wise comparisons were made between the (C16:0 + C17:0cpa)/(C18:1 + C19:0cpa), as these fatty acids dominated the membrane composition under each growth condition examined.
3. Results and discussion Alterations in the membrane fatty acid or phospholipid composition are common adaptation mechanisms for many bacteria in response to environmental stress. The cariogenic organism S. mutans alters its membrane fatty acid profile in response to acidification of its environment, specifically by increasing the proportions of monounsaturated membrane fatty acids when grown in low pH environments [3,21]. If the ability to increase or produce mono-unsaturated fatty acids is prevented, the organism is rendered acid sensitive [3,24]. Since membrane fatty acid adaptation is a protective measure utilized by S. mutans, it is possible that other oral bacteria utilize this mechanism to protect against acidification of the environment. We grew four oral bacteria, both aciduric organisms and those considered acid-sensitive, in glucose-limited chemostats under controlled pH conditions. Cells were harvested and membrane fatty acids were determined by GC-FAME by Avanti Polar Lipids, Inc. Although S. sanguis has demonstrated a weak ability to adapt to acidification [17,20,25,26], it is not considered aciduric and is not as capable as S. mutans in survival of low pH environments [15,18,19,27,28]. Specifically, studies have shown that S. sanguis 10904 possesses an F-ATPase, a necessary component for low pH survival of oral streptococci, which is not as active at pH values as low as those reported for the S. mutans enzyme [13,15,18–20]. Since it is not considered aciduric and its F-ATPase does not function at pH values as low as its aciduric counterpart in S. mutans, we investigated whether the membrane fatty acid profile of S. sanguis would alter as a response to acidification.
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As shown, the membrane fatty acid content primarily consisted of C16:0, C16:1, and C18:1 (Table 1), and did not change significantly in response to pH. However, when grown in batch cultures, there were significant increases in the proportion of C18:1, similar to those reported for S. mutans [21,24]. In addition, the batch culture pH of S. sanguis was approximately 5.5, lower than the pH for chemostat growth. It may be possible that S. sanguis is capable of membrane fatty acid alterations when the environmental pH is lowered to a critical value. S. gordonii, a member of the sanguis group associated with bacterial endocarditis undergoes phenotypic acid adaptation, but is typically not considered aciduric or cariogenic [14,20,26,29]. As shown in Table 1, the organism increased the proportion of the membrane fatty acid C18:1 when the culture pH was lowered from a value of 7 to 6. Although we were unable to grow the organism in steady-state at pH 5, the pH of the batch cultures was approximately 4.7 after 16 h of growth (data not shown). The batch cultures showed even higher levels of C18:1 than those observed in the pH 6 steady-state cultures. Again, it is possible that these elevated levels are due to further acid adaptation of the batch culture than the chemostat culture. It has been shown that particular strains of oral streptococci can undergo phenotypic acid adaptation when grown with excess glucose, similar to the batch conditions used here [17]. Additionally, although both S. gordonii and S. sanguis are not considered aciduric, it is interesting to note that S. gordonii altered its membrane fatty acid profile in response to growth at pH 6 in steady-
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state, unlike S. sanguis. Although the strains were grown in different media, neither media contained detectable levels of fatty acids (data not shown). Thus, there appears to be a difference in the regulation of the fatty acid composition between these two strains. S. salivarius is considered aciduric and can rapidly adapt to acidification of its environment [13,26]. The membrane fatty acid composition of pH 7-grown S. salivarius consisted primarily of C14:0 and C16:0 fatty acids similar to S. mutans [21] (72% of the total membrane fatty acid composition in S. salivarius, 62% of the total in S. mutans). When grown at pH 5, the levels of C14:0 and C16:0 decreased dramatically to comprise only 38.6% of the total composition (Table 1); there was a simultaneous increase in the levels of C18:1 and C20:1, similar to what was reported in S. mutans [21]. The batch-grown cells had membrane fatty acid profiles similar to the pH 5-grown chemostat cells and to reports published previously [4,6–9]. Thus S. salivarius, like S. mutans, increased the proportion of mono-unsaturated membrane fatty acids in response to growth under low pH [21]. Previously, we have demonstrated that if increases in the mono-unsaturated fatty acid composition in S. mutans are blocked, the organism is rendered highly acid-sensitive [3]. It is highly possible that these increases in mono-unsaturated membrane fatty acids are necessary for S. salivarius to survive the acidic environment of dental plaque. L. casei is considered to be extremely aciduric [13,15,19,26,30], and previous reports have demonstrated that Lactobacillus species produce membranes
Table 1 Membrane fatty acid composition of S. sanguis 1 0904, S. gordonii DL 1, S. salivarius 57.1 grown at steady-state in a chemostata Fatty acid
% of total membrane fatty acid composition S. sanguis 10904
C14:0 C14:1 C16:0 C16:1 C18:0 C18:1 C20:0 C20:1 Others C14:0 + C16:0 C18:1 + C20:1 Ratioc
S. gordonii DL 1 b
S salivarius 57.1 b
pH 7
pH 6
Batch (5.5)
pH 7
pH 6
Batch (4.7)
pH 7
pH 5
Batch (4.3)b
6.1 ± 10.1 1.4 ± 0.0 18.1 ± 0.8 27.7 ± 0.6 2.2 ± 0.1 33.6 ± 1.8 ND ND 11.0 ± 2.0 24.2 ± 0.7 33.6 ± 1.8 0.7 ± 0.0
4.8 ± 0.0* 1.6 ± 0.3 14.8 ± 1.1 28.9 ± 1.7 2.3 ± 0.6 33.5 ± 0.1 ND ND 14.1 ± 0.1 19.6 ± 1.1* 33.5 ± 1.1 0.6 ± 0.0*
3.9 ± 0.1* 0.7 ± 0.4 17.6 ± 0.6 23.1 ± 2.1 4.2 ± 0.2* 43.6 ± 2.2* ND 0.1 ± 0.1 7.1 ± 3.0 21.5 ± 0.8* 43.7 ± 2.3* 0.5 ± 0.0*
21.3 ± 0.1 1.4 ± 0.0 44.3 ± 0.1 12.1 ± 0.1 3.7 ± 0.1 11.3 ± 0.1 ND ND 6.1 ± 0.2 65.5 ± 0.1 11.3 ± 0.4 5.8 ± 0.1
15.8 ± 1.5* 1.1 ± 0.0 41.5 ± 0.8* 13.2 ± 0.6 4.9 ± 0.4 19.4 ± 1.6* ND ND 4.3 ± 0.4 57.2 ± 2.3* 19.4 ± 1.6* 3.0 ± 0. 4*
4.6 ± 0.1* 0.7 ± 0.1 29.9 ± 1.4* 6.2 ± 0.6* 16.1 ± 0.9* 40.1 ± 1.8* 0.2 ± 0.1 0.2 ± 0.1 2.3 ± 0.2 34.5 ± 1.5* 40.3 ± 1.9* 0.9 ± 0.1*
20.2 ± 4.2 ND 52.3 ± 1.2 8.7 ± 0.3 5.7 ± 0.9 8.8 ± 2.2 0.1 ± 0.1 1.5 ± 0.8 2.9 ± 0.4 72.2 ± 3.0 10.2 ± 3.0 7.5 ± 2.5
6.3 ± 3.8 ND 32.3 ± 2.0* 4.3 ± 2.0 12.6 ± 5.6 23.1 ± 0.9* 3.3 ± 0.2* 16.0 ± 2.1* 2.3 ± 0.6 38.6 ± 5.8* 39.1 ± 3.0* 1.0 ± 0.2
5.9 ± 0.3* ND 37.6 ± 0.1* 6.6 ± 1.2 7.6 ± 0.4 28.2 ± 1.6* 0.6 ± 0.0* 12.4 ± 0.8* 1.3 ± 0.4 43. 5 ± 0.4* 40.6 ± 2.3* 1.1 ± 0.1
ND, not detected. Average % fatty acid ± standard deviation. n = 2. a Each strain was grown in a chemostat at least two times at a dilution rate of 0.24 h 1. Aliquots were collected after 10 generations at each pH value. b Batch cultures were grown overnight at least two times in an environment enriched with 5% CO2. pH of the batch cultures after 16 h of growth is indicated. c Ratio is (C14:0 + C16:0)/(C18:1 + C20:1). * Statistically significant difference between that condition and pH 7 value, p < 0.05.
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Table 2 Membrane fatty acid composition of L. casei 4646a Fatty acid
C14:0 C14:1 C16:0 C16:1 C17:0 cpa C18:0 C18:1 C19:0 cpa C20:0 C20:1 Others C16:0 ± C17:0 cpa C18:1 ± Cl9;0 cpa Ratioc
% of total membrane fatty acid composition pH 7
pH 5
Batch(3.8)b
3.0 ± 0.1 0.3 ± 0.1 42.7 ± 0.1 4.8 ± 0.4 21.4 ± 0.4 0.9 ± 0.0 11.8 ± 1.3 10.2 ± 0.8 0.0 ± 0.0 0.0 ± 0.0 5.0 ± 0.6 64.1 ± 0.6 22.0 ± 0.5 2.9 ± 0.1
3.7 ± 1.1 0.3 ± 0.1 30.4 ± 2.6* 7.9 ± 2.3 2.7 ± 0.3* 2.6 ± 0.8 25.3 ± 0.1* 25.1 ± 0.6* 0.0 ± 0.0 0.0 ± 0.0 2.0 ± 0.4 33.1 ± 2.3* 50.3 ± 0.6* 0.7 ± 0.0*
4.4 ± 0.6 0.6 ± 0.8 21.2 ± 8.5 16.2 ± 4.6 0.0 ± 0.0* 1.7 ± 0.2 25.9 ± 2.9* 18.6 ± 3.4 0.0 ± 0.0 0.0 ± 0.0 11.5 ± 4.2 21.2 ± 8.5* 44.5 ± 0.5* 0.5 ± 0.2*
ND, Not Detected Average % fatty acid ± standard deviation, n = 2. a L casei was grown in a chemostat two times at a dilution rate of 0.1 h 1. Aliquots were collected after 10 generations at each pH value. b Batch cultures were grown two times in an environment enriched with 5% CO7. Indicated is the pH of the cultures after 30 h of growth. c Ratio is (C16:0 + C17:0cpa)/ (C18:1 + C19:0cpa). * Statistically significant between that condition and pH 7 value, p < 0.05.
that contain cyclopropane fatty acids [5,10]. We examined the membrane fatty acid content of L. casei 4646 after growth at neutral and acidic pH values to determine whether the aciduric bacterium altered its membrane fatty acid content. As shown in Table 2, L. casei does alter its fatty acid content in response to acidification. L. casei, similar to S. mutans, S. gordonii, and S. salivarius increased the proportion of C18:1 in response to acidification, again suggesting that increased levels of longchained, monounsaturated fatty acids are necessary for low pH survival and is a mechanism commonly used by bacteria. In addition, L. casei increased the proportion of cyclopropane fatty acid C19:0 with a concomitant decrease in the levels of C16:0 and cyclopropane fatty acid C17:0. While the relative total amounts of cyclopropane fatty acids did not change with respect to growth pH, the proportion of long versus shorter chained cyclopropane fatty acids did alter. It appears that increased fatty acid length is another important membrane alteration to increase survival in acidic environments. The necessity for cyclopropane fatty acids in low pH survival of E. coli has been demonstrated, and it is likely that they are significant for the survival of L. casei as well [22,31]. In addition, cyclopropane fatty acids have also been associated with disease persistence in Mycobacterium tuberculosis and with the infectious dose of E. coli O157:H7 [32,33]. L. casei is considered to be more aciduric than S. mutans and the other oral streptococci, so the presence of cyclopropane fatty acids may increase the acid tolerance of the lactobacilli, although this remains to be determined. We have demonstrated that the oral bacteria S. gordonii. S. salivarius, and L. casei undergo membrane fatty acid alterations as a result of environmental acidification.
This may be an additional mechanism utilized by the bacteria to increase their survival during the repeated cycles of acidification in dental plaque. Recent work with S. mutans suggest that monounsaturated membrane fatty acids are necessary for maintenance of DpH across the membrane, and it is likely that this function is conserved among the species examined here [24]. It seems likely that other bacteria that must survive pH fluctuations undergo similar membrane changes. Membrane fatty acid alterations may represent a generalized response utilized by many bacteria to survive environmental stress. New therapeutics designed to target the mechanisms responsible for membrane fatty acid adaptation could increase the current arsenal employed to combat disease.
Acknowledgements The authors wish to thank Robert Marquis for helpful discussion throughout. We also thank Roberta Faustoferri for technical expertise. This work was supported by grants from the NIH, National Institute for Dental and Craniofacial Research DE-11549 and DE-06127. E.M.F and J.K.K. were supported by the Rochester Training Program in Oral Infectious Diseases, T32-DE07165.
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