Metabolic activity of probiotics—Oxalate degradation

Metabolic activity of probiotics – oxalate degradation C. Murphy, S. Murphy, F. O’Brien, M. O’Donoghue, T. Boileau, G. Sunvold, G. Reinhart, B. Kiely, F. Shanahan, L. O’Mahony To cite this version: C. Murphy, S. Murphy, F. O’Brien, M. O’Donoghue, T. Boileau, et al.. Metabolic activity of probiotics – oxalate degradation. Veterinary Microbiology, Elsevier, 2009, 136 (1-2), pp.100. ฀10.1016/j.vetmic.2008.10.005฀. ฀hal-00532521฀ HAL Id: hal-00532521 https://hal.archives-ouvertes.fr/hal-00532521 Submitted on 4 Nov 2010 HAL is a multi-disciplinary open access archive for the deposit and dissemination of scientific research documents, whether they are published or not. The documents may come from teaching and research institutions in France or abroad, or from public or private research centers. L’archive ouverte pluridisciplinaire HAL, est destinée au dépôt et à la diffusion de documents scientifiques de niveau recherche, publiés ou non, émanant des établissements d’enseignement et de recherche français ou étrangers, des laboratoires publics ou privés. Accepted Manuscript Title: Metabolic activity of probiotics – oxalate degradation Authors: C. Murphy, S. Murphy, F. O’Brien, M. O’Donoghue, T. Boileau, G. Sunvold, G. Reinhart, B. Kiely, F. Shanahan, L. O’Mahony PII: DOI: Reference: S0378-1135(08)00481-1 doi:10.1016/j.vetmic.2008.10.005 VETMIC 4234 To appear in: VETMIC Received date: Revised date: Accepted date: 16-5-2008 2-10-2008 6-10-2008 Please cite this article as: Murphy, C., Murphy, S., O’Brien, F., O’Donoghue, M., Boileau, T., Sunvold, G., Reinhart, G., Kiely, B., Shanahan, F., O’Mahony, L., Metabolic activity of probiotics – oxalate degradation, Veterinary Microbiology (2008), doi:10.1016/j.vetmic.2008.10.005 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. Manuscript Metabolic activity of probiotics – oxalate degradation 1 2 C. Murphy1, S. Murphy1, F. O’Brien1, M. O’Donoghue2, T. Boileau3, G. Sunvold3, G. 4 Reinhart3, B. Kiely1, F. Shanahan4, L. O’Mahony1,4. us cr ip 5 1 6 2 7 Alimentary Health Ltd., National University of Ireland, Cork, Ireland. Department of Microbiology, National University of Ireland, Cork, Ireland. 3 8 4 Procter & Gamble Pet Health and Nutrition, Ohio, US. Alimentary Pharmabiotic Centre, National University of Ireland, Cork, Ireland. an 9 t 3 12 M 10 13 Keywords: Hyperoxaluria, Oxalic acid, Probiotics te d 11 14 ep 15 Author for correspondence: 17 Dr. L. O’Mahony 18 Alimentary Pharmabiotic Centre, 19 National University of Ireland, Cork, 20 Cork. 21 Telephone: +353 (0) 21 4901372 22 Fax: +353 (0) 21 4276318 23 E-mail: [email protected] Ac c 16 1 Page 1 of 33 24 25 Abstract Urinary tract stones are an important clinical problem in human and veterinary medicine. Hyperoxaluria is the single strongest promoter of kidney stone formation. The 27 aims of the present study were to, (a) evaluate oxalate degradation by a range of 28 Bifidobacteria species and Lactobacillus species isolated from the canine and feline 29 gastrointestinal tract in vitro and, (b) to determine the impact of oxalate degradation by 30 selected strains in vivo. The bacteria were grown in oxalate-containing media and their 31 ability to degrade oxalate in vitro was determined using reverse-phased HPLC. 32 Bifidobacteria species and Lactobacillus species that degraded oxalate in vitro and 33 survived gastric transit were selected for further examination. The selected probiotics 34 were fed to rats for 4 weeks. Urine was collected at week’s 0, 2 and 4 and oxalate levels 35 determined by HPLC. In vitro degradation was detected for 11/18 of the Lactobacillus 36 species. In contrast, the capacity to degrade oxalate was not detected for any of the 13 37 Bifidobacterium species tested. Lactobacillus animalis 223C, Lactobacillus murinus 38 1222, Lactobacillus animalis 5323 and Lactobacillus murinus 3133 were selected for 39 further investigation in a rat model. Urinary oxalate levels were significantly reduced 40 (p<0.05) in animals fed L. animalis 5323 and L. animalis 223C but were unaltered when 41 fed L. murinus 1222, L. murinus 3133 or placebo. Probiotic organisms vary widely in 42 their capacity to degrade oxalate. In vitro degradation does not uniformly translate to an 43 impact in vivo. The results have therapeutic implications and may influence the choice of 44 probiotic, particularly in the setting of enteric hyperoxaluria. Ac c ep te d M an us cr ip t 26 2 Page 2 of 33 45 1. Introduction Hyperoxaluria complicated by renal tract stones is an important clinical problem 47 in humans, particularly those with enteric hyperoxaluria secondary to conditions such as 48 Crohn’s disease (Kumar et al., 2004). In veterinary medicine, domestic animals, such as 49 cats and dogs, are particularly prone to oxalate stones. Currently, there is no successful 50 medical dissolution protocol, and renal stones must be removed or disrupted by physical 51 methods. Epidemiological studies over the last decade have associated a decrease in 52 struvite calculi with an increase in calcium oxalate renal stone formation (Hesse et al., 53 1998; Lekcharoensuk et al., 2001). Acidification of commercial diets to maintain urine 54 pH between 6.0 and 6.4 reduces struvite crystal formation but increases the risk of 55 calcium oxalate formation in companion animals (Buffington and Chew, 1996).Oxalic 56 acid and its salts are widely distributed in dry commercially prepared dog food 57 (Hodgkinson, 1977; Stevenson et al., 2003). Increased dietary oxalate results in increased 58 urinary oxalate and calcium oxalate relative supersaturation in healthy adult dogs 59 (Stevenson et al., 2003). us cr ip an M d te ep 60 t 46 While some components of the enteric bacterial flora, (such as Oxalobacter formigenes) have oxalate degrading capacity, these organisms are not uniformly present 62 in all animals (Allison et al., 1986; Sidhu et al., 2001). However, dietary 63 supplementation with probiotics has emerged as a potential strategy for increasing the 64 degradation of dietary oxalate (Campieri et al., 2001; Weese et al., 2004). Therefore, the 65 purpose of our study was to screen a range of Lactobacillus species and Bifidobacteria 66 species derived from the feline and canine gastrointestinal tract for oxalate degradation 67 capacity in vitro and then to determine the impact of feeding such strains on urinary 68 oxalate excretion in vivo. Ac c 61 3 Page 3 of 33 69 70 2. Materials and methods 71 2.1 Probiotic stain isolation The small intestine, caecum or colon of cats and dogs were dissected post mortem 73 and the removed tissue washed in Ringers solution (Oxoid, Basingstoke, Hampshire, UK) 74 to remove loosely adherent bacteria. The tissue was vortexed and homogenised in 75 Ringers solution to select adherent bacteria. The supernatants from the wash and vortex 76 steps were plated on de Man, Rogosa, Sharpe (MRS) agar (Oxoid, Basingstoke, 77 Hampshire, UK) supplemented with 20 g/ml vancomycin (Sigma-Aldrich Chemie, St. 78 Louis, MO, USA) and Wilkins Chalgren Agar (Oxoid, Basingstoke, Hampshire, UK) 79 supplemented with 50 g/ml mupirocin (Oxoid, Basingstoke, Hampshire, UK) for 80 Lactobacillus species and Bifidobacteria species, respectively. The plates were incubated 81 at 37C in an anaerobic environment for 72 h. Isolated colonies were re-streaked to 82 ensure purity. Isolates from MRS agar + vancomycin plates were re-streaked on MRS 83 agar and isolates from Wilkins Chalgren Agar + mupirocin were re-streaked on 84 Reinforced Clostridia Agar (RCA: Oxoid, Basingstoke, Hampshire, UK) supplemented 85 with 0.05% (v/v) L-cysteine hydrochloride (Sigma-Aldrich Chemie, St. Louis, MO, 86 USA) for the purification of Lactobacillus species and Bifidobacteria species, 87 respectively. Following purification, single strain cultures were identified on the basis of 88 colony morphology, gram reaction, catalase activity and the Fructose-6-phosphate 89 phosphoketolase assay. Gram-positive, catalase negative rods were genetically 90 characterised using primers specific for the 16 S intergenic spacer region and 91 Lactobacillus species and Bifidobacteria species isolates were further examined. Ac c ep te d M an us cr ip t 72 4 Page 4 of 33 Lactobacillus species strains were routinely cultured in MRS broth at 37C in an 93 anaerobic environment for 24 h. Bifidobacteria species isolates were routinely cultured in 94 MRS broth supplemented with 0.05% (v/v) L-cysteine hydrochloride and incubated at 95 37C in an anaerobic environment for 48 h. Lactobacillus species and Bifidobacteria 96 species stocks are maintained in 40% glycerol at -80º C (Alimentary Health Ltd., 97 National University of Ireland, Cork, Ireland). 98 99 us cr ip t 92 2.2 Assaying Lactobacillus and Bifidobacteria isolates for growth in ammonium oxalate media and determining oxalate-degrading capability 101 The procedure for the determination of oxalate-degrading capacity of probiotic isolates 102 was based on the method previously described by Campieri et al. (2001). Briefly, 5 ml of 103 filtered sterilised ammonium oxalate solution [20 mM/l ammonium oxalate and 40 g/l 104 dextrose (Roqette, Lestrem, France)] was added to 5 ml of base media (Protease peptone 105 20 g/l, yeast extract 10 g/l, Tween 80 2 ml/l, KH2PO4 4 g/l, NA acetate 10 g/l, di- 106 Ammonium-hydrogen-citrate 4 g/l, MgSO4.7H2O 0.1 g/l and MnSO4 0.1 g/l). All reagents 107 were supplied by either Sigma-Aldrich (St. Louis, MO, USA) or BDH Laboratory 108 supplies, Poole, UK; unless otherwise stated. Culture broths were inoculated at 2% into 109 base media and base media containing 20 mM ammonium oxalate. The base media was 110 supplemented with 0.05% (v/v) L-cysteine hydrochloride when inoculating 111 Bifidobacteria species and all cultures were incubated anaerobically at 37C for 48 h. A 112 media control (ammonium oxalate base media) was prepared as above, but without the 113 inoculation of bacteria. Optical density (600 nm) and plate counts (colony forming 114 units/ml) were performed to determine growth of each strain in ammonium oxalate base Ac c ep te d M an 100 5 Page 5 of 33 media, which was compared to growth in base media. Ammonium oxalate base media 116 cultures and the media control were centrifuged at 3000 rpm for 10 min and the 117 supernatants filter sterilised using 0.45 M filters (Sartorius AG, Goettingen, Germany). 118 The culture filtrates were stored at 4C until plate counts were recorded and HPLC 119 analysis was performed on strains that grew in 20 mM ammonium oxalate base media. us cr ip t 115 120 121 122 2.3 Chemicals and materials for HPLC All chemicals were of spectral or analytical grade. Unless otherwise stated, all chemicals employed were obtained from Sigma-Aldrich (St. Louis, MO, USA) or BDH 124 Laboratory supplies, Poole, UK. HPLC grade water (Reagecon, Shannon, Ireland) was 125 utilised throughout the experiments. The procedure for the determination of oxalic acid in 126 samples by HPLC was based on the method previously described by Khaskhali et al. 127 (1996). The mobile phase was composed of 0.25% potassium dihydrogen phosphate and 128 0.0025 M tetrabutylammonium hydrogensulphate, buffered at pH 2.0 with 129 orthophosphoric acid. The mobile phase was filtered through a 0.2 m nylon membrane. 130 Aqueous oxalic acid standards were prepared in the range 0.02-20 mM. These solutions 131 were stable for 3 months at 4C. 133 134 M d te ep Ac c 132 an 123 2.4 Apparatus and chromatographic conditions Chromatographic analysis was performed using a Spectraseries 100 135 (Thermoseparation Products, Minnesota, USA) with a chromjet integrator, UV detector 136 and a Synergi Hydro-RP column, 4 m, 250 x 4.6 mm I.D. (Phenomenex, Cheshire, UK). 137 The analytic column was routinely cleaned by rinsing the column with: 94% water/5% 6 Page 6 of 33 acetonitrile, tetrahydrofluran, 95% acetronitrile/5% water and mobile phase for 20 min 139 each. The column was purged by pumping the mobile phase at 4 ml/min for 3 min and 140 equilibrated by pumping the mobile phase to waste. The detector wavelength was fixed at 141 210 nm. The total cycle time was 35 min with 20 l injections from each sample. At the 142 end of each run, acetonitrile: HPLC-grade water (65:35) was pumped through the column 143 for 15 min prior to storage. us cr ip t 138 144 145 2.5 Preparation of filtrate samples 20 mM, 15 mM, 10 mM, 5 mM and 2 mM ammonium oxalate standards were 147 prepared from 200 mM ammonium oxalate stock solution. All filtrates and standards 148 were diluted 1:50 in mobile phase and analysed using HPLC. M an 146 149 2.6 Survival in a low pH environment. d 150 Probiotic strains must be capable of resisting the effects of a low pH environment. 152 Bacterial cells were harvested from overnight cultures, washed twice in phosphate buffer 153 (pH 6.5) and resuspended in the MRS broth adjusted with 1 N HCl to pH 2.5. The cells 154 were incubated anaerobically @ 37C and their survival measured at intervals of 0, 30, 155 60, 120, 180, 240 and 360 min using the plate count method. 157 158 ep Ac c 156 te 151 2.7 Resistance to bile salts Resistance to bile was examined using MRS agar plates supplemented with 0.5, 159 1.0 and 5.0 % (w/v) porcine bile (Sigma-Aldrich Chemie, St. Louis, MO, USA). 160 Lactobacillus species probiotics were inoculated into MRS broth and incubated at 37C 7 Page 7 of 33 under anaerobic conditions for 24 h. Strains were spot inoculated (10µl) onto the various 162 concentrations of porcine bile plates and incubated at 37C under anaerobic conditions 163 for 48 h. The growth rate on porcine bile plates were compared to the growth rate on 164 MRS agar plates and recorded. 165 166 2.8 Tolerance to freeze drying process and stability us cr ip t 161 The probiotic strains were grown overnight in MRS broth, centrifuged and 168 resuspended in cryoprotectant (18% reconstituted skim milk, 2 % sucrose). The mixtures 169 were then frozen at -20C for 24 Hrs and then freeze dried for another 24 Hrs. The 170 mixtures were freeze-dried at a vacuum pressure of 133 x 10-3 mBar with a condenser 171 temperature of -53C. All strains were examined for stability to freeze-drying and their 172 shelf life at room temperature was assessed for one month post-processing by MRS plate 173 counting techniques. te 174 d M an 167 2.9 Generation of spontaneous rifampicin-resistant variants of isolated probiotics 176 Selected probiotics were streaked onto MRS agar for Lactobacillus species 177 isolates and RCA supplemented with 0.05% L-cysteine hydrochloride for Bifidobacteria 178 species isolates. All isolates were incubated at 37C in an anaerobic environment for 48 179 h. Isolates were sub-cultured onto appropriate agar plates containing 100 g/ml 180 rifampicin and incubated at 37C in an anaerobic environment for 72 h. Spontaneous 181 rifampicin resistant variants (RifR) were stocked in 40% glycerol (Sigma-Aldrich 182 Chemie, St. Louis, MO, USA), stored at -80 C and checked for their continuous 183 resistance to 100 g/ml rifampicin by restreaking onto appropriate agar plates containing Ac c ep 175 8 Page 8 of 33 184 100 g/ml rifampicin and incubated at 37C in an anaerobic environment for 48 h. 185 Growth curves of isolates and RifR isolates were performed to ensure the growth rate was 186 not altered. 188 189 2.10 In vivo gastric transit of selected probiotic isolates us cr ip t 187 15 female Spague-Dawley rats of similar age and weight were enrolled in the study. Freeze dried RifR probiotic powders were resuspended in an appropriate volume of 191 water to ensure a does of ~ 9.8 x 109 colony-forming units (cfu) for L. animalis 223C, L. 192 murinus 1222, L. animalis 5323 and L. murinus 3133 or 0 cfu control freeze dried product 193 for the placebo group. The resuspended powders were administered, ad libitum, for 6 194 days (n=3 animals per group). Rats were weighted daily and the volume of probiotic 195 consumed was calculated daily. Rat faecal pellets were collected prior to feeding (Day 0) 196 and on Days 1, 3 and 6 (post probiotic feeding). All faecal pellets were weighed and 197 resuspended in 1 ml Ringers (Oxoid, Basingstoke, Hampshire, UK). The colony forming 198 units/g was determined by plating onto MRS agar containing 100 g/ml rifampicin, in 199 order to facilitate uncomplicated identification of the freeze dried RifR probiotics from all 200 other Lactobacilli. 202 203 M d te ep Ac c 201 an 190 2.11 In vivo urinary oxalate levels using selected probiotics 30 female Sprague-Dawley rats of similar age and weight were enrolled in the 204 study. Freeze dried probiotic powders were resuspended in an appropriate volume of 205 water to ensure a does of ~ 2 x 109 cfu for L. animalis 223C, L. murinus 1222, L. animalis 206 5323 and L. murinus 3133 or 0 cfu control freeze dried product for the placebo group. 9 Page 9 of 33 207 The resuspended powders were administered, ad libitum, for 4 weeks (n=6 animals per 208 group) Rats were weighed weekly and urine samples were obtained on Weeks 0, 2 and 4 209 by placing the animals in metabolic cages for a 24 h period. 212 2.12 Preparation of urine samples us cr ip 211 t 210 10 ml of a 24 hour sample was obtained from the metabolic cage and placed in polyethylene bottles to which 10 ml of 6  hydrochloric acid was added as a 214 preservative. Deproteinisation of the samples was performed at ambient temperature by 215 mixing a homogeneous urine sample (10 ml) from each collection with 0.5 g crystalline 216 sulfosalicylic acid and after 10 min filtering the mixture through a 0.45 m Minisart filter 217 (Khaskhali et al., 1996). M an 213 220 2.13 Statistical analysis Statistical analysis of the in vitro results was performed using a paired student t- te 219 d 218 tests. Changes in rat urinary oxalate excretion levels over time were assessed using a one- 222 way analysis of variance (ANOVA) with replicates. Ac c ep 221 10 Page 10 of 33 223 3. Results 224 3.1 In vitro growth and oxalate degradation by probiotics of canine and feline origin. Thirteen Bifidobacteria species and 18 Lactobacillus species were included in the 226 in vitro assessment, which were identified using 16S intergenic spacer sequencing. These 227 strains included 11 B. longum strains (feline-derived), 1 B. globosum strain (canine- 228 derived), 1 B. animalis strain (canine-derived), 1 L. acidophilus strain (feline- 229 derived), 5 L. reuteri strains (feline-derived), 8 L. animalis strains (7 canine-derived 230 & 1 feline-derived), 1 L. salivarius strain (canine-derived) and 3 L. murinus strains 231 (canine-derived). All selected isolates grew in the presence of 20 mM ammonium 232 oxalate illustrating that oxalate at this concentration is not toxic to LAB. The average 233 cfu/ml of isolates, grown in the presence of 20 mM ammonium oxalate, was 2.3 x 108 234 cfu/ml. This was comparable to growth of isolates in base media. Supernatants from 235 isolates were subsequently analysed using HPLC. A media control (base media + 20 mM 236 ammonium oxalate) was included in order to provide a 20 mM ammonium oxalate 237 standard. ep te d M an us cr ip t 225 The ability of Lactic Acid Bacteria (LAB) to degrade oxalate was strain 239 dependant. No oxalate degradation was detected for any of the Bifidobacterium species 240 isolates when compared to the 20 mM ammonium oxalate media control (Fig. 1). 241 Oxalate degradation was detected for 11/18 (61%) of the Lactobacillus species when 242 compared to the ammonium oxalate media control (Fig. 2). L. acidophilus, L. reuteri and 243 L. salivarius isolates did not demonstrate oxalate degradation, but L. animalis and L. 244 murinus isolates demonstrated significant oxalate degradation. Two representative 245 isolates from L. animalis and two representative isolates from the L. murinus group were 246 selected for further examination in an in vivo rat model. Mean rate of in vitro oxalate Ac c 238 11 Page 11 of 33 247 degradation for the selected strains was 0.15 mM/h (L. animalis 223C – feline isolate), 248 0.15 mM/h (L. murinus 1222 – canine isolate), 0.14 mM/h (L. animalis 5323 – canine 249 isolate) and 0.09 mM/h (L. murinus 3133 – canine isolate). 252 3.2 Assessment of gastric transit of probiotic bacteria in vitro us cr ip 251 t 250 Prior to reaching the intestinal tract, probiotic bacteria must first survive transit through the stomach, which involves survival to stomach and bile acids. The survival of 254 selected strains to a low pH environment was assessed by adding approximately 108 255 cfu/ml of L. animalis 223C, L. murinus 1222, L. animalis 5323 and L. murinus 3133 to 256 acidified MRS broth, pH 2.5. The results indicate that all selected probiotic strains have 257 the potential to successfully transit the human stomach, as strains were viable after 360 258 minutes in a low pH environment and the loss of viability was <1.5 logs (Fig 3). M The survival of probiotic strains upon exposure to deconjugated porcine bile was d 259 an 253 examined using MRS agar plates supplemented with various concentrations of bile. All 261 selected strains survive up to 5.0 % bile acid (Table 1). 264 ep 263 3.3 Stability of bacterial strains following the freeze-drying process Ac c 262 te 260 The putative probiotic strains were examined for their stability, following the 265 freeze-drying process, for 1 month at room temperature. L. animalis 223C, L. murinus 266 1222, L. animalis 5323 and L. murinus 3133 remained at high numbers post freeze-drying 267 and demonstrated no loss of activity during storage at room temperature (Fig 4). 268 269 3.4 In vivo gastric transit of selected probiotic isolates 12 Page 12 of 33 270 Changes in rat weight were monitored daily during the gastric transit feeding trial. No significant changes in body weight were detected for the duration of the trail. The 272 volume of RifR probiotic consumed ad libitum was recorded and the dose of RifR 273 probiotic consumed was calculated based on the dose of freeze-dried probiotic supplied 274 (Table 2). The average dose of probiotic consumed/day was 9.8 x 109 CFU. The 275 consumed probiotics survived gastric transit in this rat model (Fig 5). Prior to feeding 276 probiotics (Day 0), no RifR probiotics were detected on culture plates. This baseline 277 ensures the selectively of the agar plates containing 100 µg/ml rifampicin. The RifR 278 probitics were detected in faeces from all mice in the probiotic group within 1 day of 279 feeding. During the 6 day feeding study, the RifR probiotics were recovered at 280 approximately 4.6 x 109 bacteria per gram of faeces. RifR probiotics were not cultivated 281 from any of the rats in the placebo group. The amount of RifR probiotic consumed/day is 282 equivalent to the gastric transit of the probiotics/day. No significant difference was 283 observed between groups fed different probiotics or between transit levels on Day 1, 3 or 284 6. 287 us cr ip an M d te ep 286 3.5 In vivo oxalate degradation of selected probiotics in a rat model. Ac c 285 t 271 Sprague-Dawley rats (n=6/group) received 2 x 109 cfu/day of L. animalis 223C, L. 288 murinus 1222, L. animalis 5323 and L. murinus 3133 or placebo. During the study, 24 h 289 urine specimens were obtained on Week 0, Week 2 and Week 4 by placing the rats in 290 metabolic cages. The mean urinary output per rat was 14.3mls over the 24 hours 291 (range 10.5 – 21.2mls). Rat weights were monitored for the duration of the study, and 292 demonstrated no significant difference when compared to the placebo control (Table 3). 13 Page 13 of 33 Fig. 6 illustrates the trial results with urinary oxalate levels expressed as M 294 oxalate over a 24 hour period. Urinary oxalate levels remained constant in the first 295 group of rats (not receiving a probiotic supplement). In contrast, rats consuming the 296 probiotic strains L. animalis 223C and L. animalis 5323 had decreased urinary oxalate 297 excretion. Rats consuming L. murinus 1222 and L. murinus 3133 did not have decreased 298 urinary oxalate excretion. us cr ip t 293 299 301 4. Discussion The results of this study show that some strains of Lactobacillus but not an 300 Bifidobacteria species degrade oxalate in vitro and reduce urinary oxalate excretion in 303 vivo. Several L. animalis and L. murinus isolates degrade ammonium oxalate in vitro 304 while four strains were selected for inclusion in the animal study, 2 representatives from 305 the L. animalis group (L. animalis 223C and L. animalis 5323) and 2 representatives from 306 the L. murinus group (L. murinus 1222 and L. murinus 3133). Both L. animalis strains (L. 307 animalis 223C and L. animalis 5323) reduced oxalate excretion in rats. All 4 selected 308 strains survived gastric transit. d te ep Previous studies have demonstrated oxalate degradation by O. formigenes, a gram Ac c 309 M 302 310 negative, anaerobic bacterium that inhabits the gastrointestinal tracts of humans and 311 mammals (Allison et al., 1986; Dawson et al., 1980). The presence of O. formigenes has 312 been shown to reverse hyperoxaluria in a rat model and reduce urinary oxalate excretion 313 in humans (Duncan et al., 2002; Sidhu et al., 2001). It has been suggested that the 314 absence of O. formigenes in the gastrointestinal tract correlates with the number of 315 recurrences of oxalate stone disease (Sidhu et al., 1999). However, the establishment of 14 Page 14 of 33 O. formigenes in a rat model was transient and the faecal population of O. formigenes 317 declined below the detectable limit once rats were placed on a normal diet (Sidhu et al., 318 2001). Difficult isolation and transient colonisation of O. formigenes have resulted in 319 investigators screening for alternative oxalate-degrading bacteria in the intestine, such as 320 LAB (Campieri et al., 2001; Hokama et al., 2000; Hokama et al., 2005). P. rettgeri and 321 E. faecalis appear to have a mechanism of oxalate degradation similar to O. formigenes, 322 but they were unable to maintain their oxalate degrading ability when subcultured into 323 nutrient rich medium (Hokama et al., 2000; Hokama et al., 2005). We have shown, using 324 in vitro and in vivo models, that certain probiotics offer a therapeutic strategy to reducing 325 urinary oxalate excretion. us cr ip an All four candidate strains tested degraded oxalate in vitro, but only two of these M 326 t 316 strains degraded oxalate in vivo. It is unlikely that the inability of L. murinus 1222 and L. 328 murinus 3133 to degrade oxalate in vivo could be attributed to the physiological aspects 329 of the intestinal tract (gastric acidity, peristalis, bile acids etc.) and the anti-microbial 330 defence mechanisms (adhesion, colonisation, nutrient competition etc.), as all four strains 331 transited the gut in equivalent amounts. Rather, the L. animalis and L. murinus strains 332 may interact with the host in a strain specific manner such as that demonstrated for 333 probiotic adherence to intestinal tissue and mucus (Ouwehand et al., 1999). In addition, 334 the utilisation of oxalate as a substrate for L. murinus in vivo may not be allowable at a 335 genetic level due to phenomena such a quorum sensing. This highlights the importance of 336 carefully selecting strains using in vitro characteristics, in addition to using animal 337 models to observe the biological impact in vivo. It is unlikely that the original source 338 of these strains has a significant impact on the excretion of oxalate in the rat studies Ac c ep te d 327 15 Page 15 of 33 339 as one of the successful strains was canine-derived (L. animalis 5323) while the other 340 was feline-derived (L. animalis 223C). 341 Our results suggest considerable variability in the ability of probiotics to degrade oxalate, both in vitro and in vivo. We detected oxalate degradation for 61% of the 343 Lactobacillus species examined in vitro. In contrast, Bifidobacterium species appears not 344 to possess the mechanism of oxalate degradation demonstrated by Lactobacillus spp 345 when examined in vitro. Weese et al. (2004) also reported considerable variation in 346 oxalate degradation by different probiotics in vitro. They reported a mean oxalate 347 degradation of 17.7 % for 37 LAB, but they did not further identify the strains. Campieri 348 et al. (2001) previously reported variable in vitro oxalate degradation with L. acidophilus, 349 L. plantarum, L. brevis, Streptococcus thermophilus and B. infantis. They demonstrated 350 little or no oxalate degradation in L. plantarum and L. brevis, but L. acidophilus, S. 351 thermophilus and B. infantis degraded oxalate. However, the level of in vitro oxalate 352 degradation was low, with degradation of 5.26% of 10 mM/l ammonium oxalate and 353 2.18% of 20 mM/l ammonium oxalate and in vivo degradation was assessed in a mixture 354 of freeze-dried LAB (L. acidophilus, L. plantarum, L. brevis, S. thermophilus, B. 355 infantis). Why only some probiotics strains degrade oxalate remains unclear, fuelling a 356 desire to better understand the mechanism of oxalate degradation in probiotics. O. 357 formigenes has two oxalate degrading enzymes, oxalyl-coenzyme A decarboxylase (65 358 kDa) and formyl-coenzyme A transferase (48 kDa) (Kodoma et al., 2002). While these 359 oxalate degrading enzymes have been found in Providencia rettgeri and Enterococcus 360 faecalis, it is unknown if these enzymes have been found in LAB (Hokama et al. 2005; 361 Hokama et al. 2000). Ac c ep te d M an us cr ip t 342 16 Page 16 of 33 362 The detected oxalate degradation in this study appears to be interspecies dependent, with L. animalis and L. murinus degrading oxalate in vitro and L. acidophilus, 364 L. reuteri and L. salivarius demonstrating no oxalate degradation in vitro. Indeed, only L. 365 animalis strains and not L. murinus strains degraded oxalate in vivo. Other studies have 366 demonstrated considerable interspecies variation in metabolic activity; in particular the 367 ability to produce the health-promoting fatty acid conjugated linoleic acid (CLA) from 368 free linoleic acid (Coakley et al., 2003). They demonstrated considerable interspecies 369 variation, with B. breve and B. dentium being the most efficient CLA producers. us cr ip t 363 371 an 370 5. Conclusion We have highlighted the metabolic potential of probiotics by examining one 373 specific metabolite, but mining the gut microbiota for further health promoting effects is a 374 viable option for future dietary management strategies of specific metabolic symptoms or 375 dysfunction. Future studies should also consider the development of an effective oxalate 376 degrading synbiotic (probiotic + prebiotic) by tailoring a prebiotic towards the specific 377 organism and investigating this combination using in vitro and in vivo studies (Weese et 378 al., 2004). Given that all rats tolerated the probiotic treatment well and strains L. animalis 379 223C and L. animalis 5323 in particular demonstrated superior oxalate degradative 380 capability, these strains are being further investigated as a probiotic food supplement for 381 the prevention and treatment of hyperoxaluria and renal stone formation. Ac c ep te d M 372 382 383 Acknowledgements 384 The authors are supported in part by Science Foundation Ireland in the form of a 385 centre grant (Alimentary Pharmabiotic Centre), by the Health Research Board (HRB) of 17 Page 17 of 33 386 Ireland, the Higher Education Authority (HEA) of Ireland, and the European Union 387 (PROGID QLK-2000-00563). 388 Disclosures 390 us cr ip t 389 Alimentary Health is a multi-departmental university campus-based research 391 company, which investigates host-flora interactions. The content of this article was 392 neither influenced nor constrained by this fact. an 393 394 M 395 396 References 398 Allison, M.J., Cook, H.M., Milne, D.B., Gallagher, S., Clayman, R.V., 1986, Oxalate 401 402 403 404 405 te ep 400 degradation by gastrointestinal bacteria from humans. J Nutr 116, 455-460. Buffington, C.A., Chew, D.J., 1996, Intermittent alkaline urine in a cat fed an acidifying diet. J Am Vet Med Assoc 209, 103-104. Ac c 399 d 397 Campieri, C., Campieri, M., Bertuzzi, V., Swennen, E., Matteuzzi, D., Stefoni, S., Pirovano, F., Centi, C., Ulisse, S., Famularo, G., De Simone, C., 2001, Reduction of oxaluria after an oral course of lactic acid bacteria at high concentration. Kidney Int 60, 1097-1105. 18 Page 18 of 33 406 Coakley, M., Ross, R.P., Nordgren, M., Fitzgerald, G., Devery, R., Stanton, C., 2003, 407 Conjugated linoleic acid biosynthesis by human-derived Bifidobacterium species. 408 J Appl Microbiol 94, 138-145. Dawson, K.A., Allison, M.J., Hartman, P.A., 1980, Isolation and some characteristics of t 409 anaerobic oxalate-degrading bacteria from the rumen. Appl Environ Microbiol 40, 411 833-839. 412 us cr ip 410 Duncan, S.H., Richardson, A.J., Kaul, P., Holmes, R.P., Allison, M.J., Stewart, C.S., 2002, Oxalobacter formigenes and its potential role in human health. Appl 414 Environ Microbiol 68, 3841-3847. 416 Hesse, A., Steffes, H.J., Graf, C., 1998, Pathogenic factors of urinary stone formation in animals. J Anim Phys Anim Nutr 80, 108-119. M 415 an 413 Hodgkinson, A., 1977, Oxalic acid in biology and medicine. Academic press London. 418 Hokama, S., Honma, Y., Toma, C., Ogawa, Y., 2000, Oxalate-degrading Enterococcus te 420 faecalis. Microbiol Immunol 44, 235-240. Hokama, S., Toma, C., Iwanaga, M., Morozumi, M., Sugaya, K., Ogawa, Y., 2005, ep 419 d 417 Oxalate-degrading Providencia rettgeri isolated from human stools. Int J Urol 12, 422 533-538. Ac c 421 423 Hoppe, B., von Unruh, G., Laube, N., Hesse, A., Sidhu, H., 2005, Oxalate degrading 424 bacteria: new treatment option for patients with primary and secondary 425 426 hyperoxaluria? Urol Res 33, 372-375. Khaskhali, M.H., Bhanger, M.I., Khand, F.D., 1996, Simultaneous determination of 427 oxalic and citric acids in urine by high-performance liquid chromatography. J 428 Chromatogr B Biomed Appl 675, 147-151. 19 Page 19 of 33 429 430 431 Kodama, T., Akakura, K., Mikami, K., Haruo, I., 2002, Detection and identification of oxalate-degrading bacteria in human faeces. Int J Urol 9, 392-397. Kumar, R., Ghoshal, U.C., Singh, G., Mittal, R.D., 2004, Infrequency of colonization with Oxalobacter formigenes in inflammatory bowel disease: possible role in 433 renal stone formation. J Gastroenterol Hepatol 19, 1403-1409. us cr ip 434 t 432 Lekcharoensuk, C., Osborne, C.A., Lulich, J.P., Pusoonthornthum, R., Kirk, C.A., Ulrich, L.K., Koehler, L.A., Carpenter, K.A., Swanson, L.L., 2001, Association between 436 dietary factors and calcium oxalate and magnesium ammonium phosphate 437 urolithiasis in cats. J Am Vet Med Assoc 219, 1228-1237. 438 an 435 Ouwehand, A.C., Niemi, P., Salminen, S.J., 1999, The normal faecal microflora does not affect the adhesion of probiotic bacteria in vitro. FEMS Microbiol Lett 177, 35- 440 38. Sidhu, H., Allison, M.J., Chow, J.M., Clark, A., Peck, A.B., 2001, Rapid reversal of d 441 M 439 hyperoxaluria in a rat model after probiotic administration of Oxalobacter 443 formigenes. J Urol 166, 1487-1491. 446 447 448 449 ep 445 Sidhu, H., Schmidt, M.E., Cornelius, J.G., Thamilselvan, S., Khan, S.R., Hesse, A., Peck, A.B., 1999, Direct correlation between hyperoxaluria/oxalate stone disease and Ac c 444 te 442 the absence of the gastrointestinal tract-dwelling bacterium Oxalobacter formigenes: possible prevention by gut recolonization or enzyme replacement therapy. J Am Soc Nephrol 10 Suppl 14, S334-340. Stevenson, A.E., Hynds, W.K., Markwell, P.J., 2003, The relative effects of supplemental 450 dietary calcium and oxalate on urine composition and calcium oxalate relative 451 supersaturation in healthy adult dogs. Res Vet Sci 75, 33-41. 20 Page 20 of 33 452 Weese, J.S., Weese, H.E., Yuricek, L., Rousseau, J., 2004, Oxalate degradation by 453 intestinal lactic acid bacteria in dogs and cats. Vet Microbiol 101, 161-166. 454 Ac c ep te d M an us cr ip t 455 21 Page 21 of 33 Table 1. Resistance of putative probiotic strains to porcine bile acids. Probiotic strains 457 were streaked onto MRS agar supplemented with porcine bile at 0.5, 1.0 and 5.0% (w/v). 458 Plates are incubated @ 37C under anaerobic conditions and growth was recorded after 459 24-48 h. Survival is illustrated as the mean percent of control (n=3; mean +/- SD). us cr ip t 456 460 461 Table 2. Quantity of freeze-dried probiotic consumed ad libitum/day by each group 462 (n=3). The average dose of probiotics consumed /day was 9.8 x 109 CFU. Doses are 463 illustrated as the mean dose/group +/- SD. an 464 Table 3. Animal weights for the placebo and test groups are illustrated over the 4 466 week feeding study. Body weight was not significantly influenced (compared to 467 placebo) by feeding probiotics to the animals. Results are illustrated as mean 468 (grams) per group (n=6) +/- SD. Ac c ep te d M 465 22 Page 22 of 33 Fig. 1. Lack of ammonium oxalate degradation by strains of Bifidobacterium species was 470 observed in vitro. No significant difference (p>0.05) was observed when compared to the 471 ammonium oxalate media control. The species examined were 11 B. longum, 1 B. 472 globosum and 1 B. animalis. Results are expressed as mean +/- SD. t 469 us cr ip 473 Fig. 2. Degradation of ammonium oxalate by strains of Lactobacillus species in vitro. No 475 significant difference (p>0.05) was observed for 7 of the strains (L. acidophilus,L. 476 reuteri, L. salivarius). 11/18 strains (L. animalis, L. murinus) demonstrated significant 477 oxalate degradation (p < 0.05) when compared to the ammonium oxalate media control. 478 The detected oxalate degradation appears to be species dependent, with L. animalis and L. 479 murinus degrading oxalate and L. acidophilus, L. reuteri and L. salivarius demonstrating 480 no oxalate degradation in vitro. Results are expressed as mean +/- SD. 481 *p<0.05 versus control te 482 d M an 474 Fig. 3. Survival of selected probiotics in a low pH environment. Bacterial cells 484 (approximately 108 cfu/ml) are resuspended into MRS broth adjusted with 1 N HCl to pH 485 2.5. Survival was measured at intervals of 0, 30, 60, 120, 180 and 360 min using the plate 486 count method. Results are expressed as mean +/- SD. Ac c 487 ep 483 488 Fig. 4. Stability of putative probiotic strains during storage for 1 month at room 489 temperature. Selected probiotic strains were examined for their stability to freeze-drying 490 and their shelf life at room temperature for one month was assessed following the process 23 Page 23 of 33 491 using the plate count method on MRS agar (n=2). Results are expressed as mean +/- 492 SD. 493 Fig. 5. Gastric transit of RifR freeze-dried probiotics. Freeze-dried RifR probiotics were 495 administered, ad libitum, at a dose of 9.8 x 109 CFU/dose to Sprague Dawley rats 496 (n=3/group). No RifR probiotics were detected on Day 0, which was prior to feeding and 497 confirms the selection of the RifR probiotics post feeding. RifR probiotics were detected 498 on Days, 1, 3 and 6 (post feeding) with no significant difference (p>0.05) observed 499 between groups fed probiotic or between the transit on Days 1, 3 and 6. Results are 500 expressed as mean +/- SD. M an us cr ip t 494 501 Fig. 6. Reduction of urine oxalate concentration by different strains of LAB in vivo. 503 Comparison of urine oxalate concentration (M/24 hours) of rats before (Week 0) and 504 after probiotic or placebo treatment (n=6/group) revealed that L. animalis 223C and L. 505 animalis 5323 significantly reduced oxalate concentration when compared to placebo. 506 Results are expressed as mean +/- SD. 507 *p<0.05 versus placebo te ep Ac c 508 d 502 24 Page 24 of 33 Ac ce pt ed M an us cr ip t Figure 1 Page 25 of 33 Figure 2 * * * * 0 an M Control 5333 3133 1222 122A 223C L. murinus L. salivarius Ac c ep te d L. acidophilus 6331 1221 5121 5342 1213 5241 5323 5131 5119 5310 5130 5316 5320 L. animalis * * * 4 L. reuteri * * * * t 8 us cr ip mMol Oxalate 12 Page 26 of 33 Figure 3 1.00E+10 1.00E+08 1.00E+07 1.00E+06 t 1.00E+05 1.00E+03 1.00E+02 1.00E+01 1.00E+00 0 30 60 120 Time (min) L. murinus 1222 L. animalis 5323 180 360 L. murinus 3133 ce pt ed M an L. animalis 223C us cr ip 1.00E+04 Ac Log cell numbers (CFU/ml) 1.00E+09 Page 27 of 33 Figure 4 1.00E+11 1.00E+10 1.00E+09 t 1.00E+07 us cr ip Log cell numbers (CFU/ml) 1.00E+08 1.00E+06 1.00E+05 1.00E+04 1.00E+03 1.00E+01 1.00E+00 L. animalis 223C an 1.00E+02 L. murinus 1222 L. murinus 3133 1 month post freeze dry - RT Ac ce pt ed M Post Freeze Dry L. animalis 5323 Page 28 of 33 Figure 5 1.E+08 Placebo L. animalis 223C 1.E+06 L. murinus 1222 1.E+02 1.E+00 0 1 3 6 ep te d M an Time (Days) us cr ip L. murinus 3133 t L. animalis 5323 1.E+04 Ac c Log cell numbers (CFU/g) 1.E+10 Page 29 of 33 Figure 6 8 Week 0 Week 2 6 * * 2 0 L. animalis 5323 an L. animalis 223C L. murinus 1222 L. murinus 3133 ep te d M Placebo us cr ip t 4 Ac c Urinary Oxalate Excretion M/24 Hours Week 4 Page 30 of 33 Table 1 Percentage survival in porcine bile ± standard deviation 0.5% Bile 1.0% Bile 5.0% Bile L. animalis 223C 66.6 ± 0.00 66.6 ± 0.00 49.95 ± 23.55 L. murinus 1222 100 ± 0.00 83.3 ± 23.55 66.6 ± 0.00 L. animalis 5323 100 ± 0.00 66.6 ± 0.00 49.95 ± 23.55 L. murinus 3133 66.6 ± 0.00 49.95 ± 23.55 49.95 ± 23.55 Ac c ep te d M an us cr ip t Strain ID Page 31 of 33 Table 2 Freeze dried Volume consumed ad Probiotic dose/day probiotic libitum/day (CFU, n=3) (CFU/ml) (ml) Placebo 0 19.00 ± 3.21 L. animalis 6.8 x 108 19.67 ± 1.67 3.4 x 108 20.67 ± 2.19 7.8 x 108 17.67 ± 3.71 3.1 x 108 17.67 ± 0.58 0 223C L. murinus L. animalis M 5323 L. murinus 1.29 x 1010 ± 1.43 x 10 8 an 1222 us cr ip t Group 1.38 x 1010 ± 1.61 x 108 5.48 x 109 ± 9.29 x 107 Ac c ep te d 3133 7.03 x 109 ± 2.90 x 10 9 Page 32 of 33 Table 3 Group Week 0 Week 2 Week 4 129 ± 5 140 ± 4 156 ± 4 L. animalis 223C 136 ± 9 157 ± 11 169 ± 12 L. murinus 1222 137 ± 11 147 ± 10 163 ± 13 L. animalis 5323 131 ± 6 148 ± 10 L. murinus 3133 125 ± 8 139 ± 9 us cr ip Placebo t Weight (grams) ± SD 162 ± 8 Ac c ep te d M an 155 ± 7 Page 33 of 33