Introduction

Microorganisms are ubiquitous components of the biosphere, colonizing both biotic and abiotic habitats. The human gut, in particular, hosts an incredibly complex ecosystem formed through microbial colonization. These microorganisms are essential for maintaining the host’s physiological balance through symbiotic interactions (Requena et al., 2018). The growing understanding of the complex interplay between the microbiome and host health has prompted a renewed interest in manipulating the gut microbiome for therapeutic purposes. This brings the concept of probiotics (Latin for “for life”), an old expression of the modern age, defining the bacterial association that modulates the gut microbiota and promotes health, which is rooted in the early 20th century with the works of Ilya Metchnikoff (Brunser and Gotteland, 2010). Probiotic research encompasses a multifaceted approach, such as (i) the establishment of rigorous selection criteria for probiotic strains (of human origin, able to survive under host gastrointestinal system conditions, have specific health effects when consumed, suitable for industrial scale production, able to survive under storage conditions, etc.), (ii) the isolation of elite probiotic strains (Lactic acid bacteria: Lactobacillus spp., Bifidobacterium spp., Streptococcus spp., Lactococcus lactis, and some species of Enterococcus spp.) (Argyri et al., 2013), spore-forming species: Bacillus spp. (Cutting, 2011), nonpathogen yeast: Saccharomyces cerevisiae boulardii (Vohra et al., 2016), and (iii) the elucidation of their potential health benefits at various dosage levels (Requena et al., 2018).

The COVID-19 pandemic has catalyzed more rapid growth in the global probiotic market, which was already expanding (Misra et al., 2021). Previously consumed by individuals seeking to alleviate digestive issues or adopt a healthier lifestyle, probiotics have become a first-line choice for consumers aiming to bolster their immune systems in the wake of the pandemic. Although SARS-CoV-2 is the causative agent of severe acute respiratory syndrome, it has been reported that it can also induce gastrointestinal infections (Nayebi et al., 2022). Oral probiotics have been shown to exhibit antiviral effects and improve gut health to restore homeostasis (Hung et al., 2021), leading to their recommendation as adjunctive therapy for COVID-19 (Xavier-Santos et al., 2022). In this context, while probiotics are generally thought to inhibit microbial adhesion, enhance intestinal barrier function, and strengthen the immune system (Stavropoulou and Bezirtzoglou, 2020), they are also reported to exert specific effects, including modulation of cytokine production by influencing intestinal epithelial cells, increased IgA secretion, activation of phagocytosis, modulation of regulatory T cell function, promotion of dendritic cell maturation, reinforcement of mucosal barriers, and reduction of viral entry (Bottari et al., 2021; Patra et al., 2021). These mechanisms are believed to facilitate viral clearance and prevent bacterial coinfections associated with COVID-19 (Patra et al., 2021). In addition to consumer demands, the probiotic market, restricted to a narrow product range, has experienced significant expansion during the pandemic, driven by the introduction of new product varieties and the growth of e-commerce platforms. In this context, the probiotic market projections indicate a market value of 77.09 billion USD by 2025 (Baral et al., 2021).

Consumers may prefer commercial probiotic dietary supplements over other probiotic forms because of their ease of transportation, consumption, dosage control, and longer shelf life (Zheng et al., 2017). Probiotic dietary supplements, which come in various formats such as capsules, tablets, powders, and liquids, typically contain millions to billions of commercially manufactured probiotic bacteria, often representing a diverse combination of genera and species (Meybodi et al., 2017).

While the ideal dosage of probiotics remains a subject of ongoing research, the recommended daily intake of probiotics ranges from 10⁶ to 10⁹ CFU/mL or g viable microorganisms. The optimal dosage of probiotics may fluctuate based on the specific strain, the intended health outcomes, the administration duration, the probiotic product type, and the probiotic strain’s viability (Aureli et al., 2010; Martinez et al., 2015; Mazzantini et al., 2021). On the other side, the various factors, including exposure to water, oxygen, strong acids, bile, heat, etc., during storage and oral administration, may also influence the viability and dosage levels of probiotics (Russell et al., 2011; Wang et al., 2022a). Given the strain-specific nature of probiotic efficacy and viability, rigorous identification and characterization of candidate probiotic strains is imperative (Soccol et al., 2014).

In 2017, the Council for Responsible Nutrition (CRN) and the International Probiotics Association (IPA) issued guidelines for probiotic labelling build upon the FAO/WHO guidelines (2002) by providing more specific recommendations tailored to the United States regulatory frameworks. Aiming to ensure consumers have access to accurate and informative labelling for probiotic products, both guidelines emphasize the importance of taxonomic identification, recommended serving size, health claims, storage conditions, and manufacturer contact information. CRN/IPA also introduces the concept of “quantitative declaration” (the explicit indication of the viable cell count in colony-forming units) and declaration of the total count of microorganisms for multispecies formulations (Council for Responsible Nutrition and International Probiotics Association, 2017).

For the reasons discussed above, accurate labelling of commercial probiotic products is paramount to empowering consumers to make informed choices (Korona-Glowniak et al., 2019; Martinez et al., 2015). A consensus among researchers highlights the prevalence of inaccuracies in probiotic product labelling regarding the overestimation of viable cell counts and the misidentification of probiotic strains (Lugli et al., 2019; Mazzantini et al., 2021; Morovic et al., 2016).

The escalating global prevalence of antibiotic resistance has raised concerns. In the concept of probiotics, the precautionary assessment that candidate probiotic bacteria selected for use do not harbor transferable antibiotic resistance genes is strongly emphasized by the FAO/WHO (2001). While probiotic strains of Lactobacillus and Bifidobacterium are generally regarded as genetically stable, the presence of high concentrations of probiotics in dietary supplements could create potential risks for antibiotic resistance (Zheng et al., 2017). In recent years, emerging evidence suggests that the potential for microorganisms in probiotic dietary supplements may serve as reservoirs for antibiotic resistance genes (Aziz et al., 2022; Gundogdu et al., 2023; Wong et al., 2015). This has raised significant concerns regarding the risk of horizontal gene transfer of antibiotic resistance determinants from probiotics to opportunistic pathogens within the intestinal microbiota (Aziz et al., 2022).

The primary objective of this study was to quantitatively enumerate and identify the Lactobacillus and Bifidobacterium strains present in commercially available probiotic dietary supplements widely marketed in Türkiye, and to assess previews of their resistance to clinically relevant antibiotics.

Materials and Methods

Sampling

A total of 10 samples were randomly purchased from retailers in Ankara, Türkiye. Seven different brands (A–G) of probiotic dietary supplements, marketed in sachet, capsule, tablet, and drop formulations, were subjected to analysis. Table 1 lists the product brand codes (A–G), sample numbers, types of products, and probiotic culture(s) claimed on the label. The samples, preserved in their original manufacturer-sealed packaging, were transported to the laboratory, maintaining cold chain requirements and aseptic conditions. The supplements were stored at 4°C and examined before expiration.

Microbial reference strains

To ensure the consistency and accuracy of assays, Lactiplantibacillus plantarum (formerly Lactobacillus plantarum) LMG2003, Latilactobacillus sakei (formerly Lactobacillus sakei) NCDO2714, Bifidobacterium longum, Staphylococcus aureus ATCC25923, and Escherichia coli LMG3083 (ETEC) were used as reference strains and kindly obtained from the culture collection of Microbiology Laboratory, Department Engineering, Faculty of Food Engineering, Ankara University.

Preparation of diluents

One gram or one milliliter of each probiotic dietary supplement was thoroughly suspended in 99 mL of Mitsuoka Buffer (4.5 g potassium dihydrogen phosphate, 6.0 g disodium hydrogen phosphate, 0.5 g L-cysteine HCl, 0.5 g Tween-80) and subsequently incubated for 30 minutes at 37°C. Serial dilutions were performed up to 10⁻⁷, utilizing Mitsuoka Buffer as the diluent medium (Champagne et al., 2011; Vinderola et al., 2019).

Enumeration and isolation of Lactobacillus and Bifidobacterium species

For isolation of Lactobacillus spp., 0.1 mL of each dilution was aseptically inoculated on MRS Agar (de Man Rogosa Sharpe; Merck^™, Germany) supplemented with 0.05% sistein and incubated under controlled conditions at 37°C for 72 hours. Post-incubation, the viable cell count was determined by enumerating the colonies on agar plates with a colony range of 30–300, using a Quebec colony counter. The results were expressed as colony-forming units per dose (CFU/dose). Selected five representative colonies were transferred to MRS broth (Merck^™, Germany) and incubated at 37°C for 18–24 hours (Todorov et al., 1999).

For isolation of Bifidobacterium spp., one milliliter of each dilution was inoculated to the sterile Petri dishes and poured over with melted MRS-NNLP agar supplemented with 100 µg/mL neomycin sulfate, 15 µg/mL nalidixic acid, 3 mg/mL lithium chloride, 200 µg/mL paramomycin sulfate, and 0.5 g/mL L-cysteine HCl. The plates were incubated anaerobically at 37°C for 72 hours using an anaerobic jar (GENbag anaer from bioMérieux). Post-incubation, the viable cell count was determined by enumerating the colonies on agar plates with a colony range of 30–300, using a Quebec colony counter. The results were expressed as colony-forming units per dose (CFU/dose). Selected five representative colonies were transferred to MRS+0.05% sistein broth (Merck^™, Germany) and incubated at 37°C for 18–24 hours under anaerobic conditions (Todorov et al., 1999). For each microbial group, the experiments were repeated in triplicate.

Following microscopic examination to elucidate their morphological characteristics, presumptive Lactobacillus spp. and Bifidobacterium spp. isolates were stored at −20°C using MRS broth and MRS+0.05% sistein broth, respectively. Both media contained 40% (v/v) glycerol (Merck^™, Germany) for cryoprotection.

Phenotypic and molecular identification of isolates

The identification of isolates employed a multifaceted approach, encompassing both phenotypic and genotypic characterization. Phenotypic characterization of Lactobacillus and Bifidobacterium isolates involved a preliminary evaluation, including assessment of cellular morphology, Gram staining reaction, and catalase activity, as described by Temiz (1999). Gram-positive, catalase-negative, rod-shaped isolates obtained from MRS agar supplemented with 0.05% cysteine were tentatively identified as members of the genus Lactobacillus. Gram-positive, catalase-negative isolates exhibiting a bifurcated morphology reminiscent of branching were tentatively classified as members of the genus Bifidobacterium from MRS-NNLP agar. The molecular identification of the strains was achieved by 16S rRNA gene sequencing, following the protocol detailed below.

Genomic DNA was extracted from the isolates using the Cell CV mini kit (Gene All, Catalog No: 106-101), and DNA concentration and purity were evaluated spectrophotometrically using a NanoDrop ND-2000 spectrophotometer (Thermo Fischer Scientific, Waltham, MA, USA). The resulting DNA samples were stored at −20°C. Amplification of the 16S rRNA gene region was conducted using polymerase chain reaction (PCR). The 2×PCR Master Mix (WizPure, Catalog No: 1401) was employed with an Applied Biosystems™ Pro Flex Thermal Cycler. Universal primers 27F (5’-AGAGTTTGATCMTGGCTCAG-3’) and 1492R (5’-CCGTCAATTCCTTTRAGTTT-3’) were amplified. Each 20 μL PCR reaction comprised 10 μL of 2×PCR Master Mix, 0.5 μL each of forward and reverse primers (100 pmol each), 2 μL of genomic DNA (100 ng/μL), and nuclease-free water to final volume. The PCR amplification protocol consisted of 35 cycles, each comprising the following steps: (i) the initial denaturation step at 95°C for 5 minutes to denature the double-stranded DNA template fully, (ii) the denaturation step at 95°C for 30 seconds to ensure complete denaturation of the template DNA, (iii) the annealing step at 52°C for 30 seconds to facilitate specific primer annealing to the target 16S rRNA gene region, (iv) the extension step at 72°C for 30 seconds to allow Taq polymerase to extend the annealed primers and amplify the target DNA, and (v) the final extension step at 72°C for 5 minutes to ensure complete extension of all amplicons. DNA fragments were electrophoresed in 1% (w/v) agarose gels to verify successful amplification. The O’Gene RulerTM 1000-bp DNA ladder (Fermentas, Finland) served as a size marker. To evaluate the integrity of the amplified DNA fragments, ethidium bromide staining (0.2 µg/mL) was followed by visualization under ultraviolet (UV) illumination using a Kodak Gel Logic 200 Imaging System (Kodak, USA). Subsequent purification using ExoSAP-IT Express PCR Cleanup Reagents (Thermo Fisher Scientific, Catalog No: 75001.200.UL) eliminated residual primers and unincorporated dNTPs, the purified DNA to be submitted for sequencing at Atlas Biotechnology (Ankara, Türkiye). To elucidate the taxonomic identity of the isolates, the generated 16S rRNA gene sequences were subjected to a comparative analysis against the National Center for Biotechnology Information (NCBI) 16S rRNA sequence database utilizing the Basic Local Alignment Search Tool (BLAST) program. L. plantarum LMG2003 and B. longum were employed as control strains to validate the accuracy of the identification process.

Assessment of antibiotic resistance testing

Antibiotic resistance of Lactobacillus and Bifidobacterium strains was conducted using the Kirby–Bauer disk diffusion method on Mueller–Hinton agar (Merck, Germany), as described by Bauer et al. (1966). Commercially available antibiotic discs (Oxoid, UK) were employed to assess resistance to a diverse panel of 18 commonly prescribed antibiotics, and the tests were performed in duplicate. The antibiotics employed in the study encompassed the following classes: (i) Beta-lactams: Ampicillin (10 μg/disc), methicillin (5 μg/disc), penicillin G (10 μg/disc), cefazolin (30 μg/disc), cefoxitin (30 μg/disc), (ii) Glycopeptides: Vancomycin (30 μg/disc), (iii) Aminoglycosides: Gentamicin (10 μg/disc), kanamycin (30 μg/disc), streptomycin (10 μg/disc), (iv) Phenicols: Chloramphenicol (30 μg/disc), (v) Lincosamides: Clindamycin (2 μg/disc), (vi) Macrolides: Erythromycin (15 μg/disc), (vii) Tetracyclines: Tetracycline (30 μg/disc), (viii) Fluoroquinolones: Enrofloxacin (5 μg/disc), Ciprofloxacin (5 μg/disc), Norfloxacin (10 μg/disc), (ix) Rifamycins: Rifampin (5 μg/disc), and (x) Trimethoprim: Trimethoprim (5 μg/disc). The plates with antibiotic discs were incubated at room temperature for at least 20 minutes prior to incubation at 37°C for 24–48 hours. Following incubation, the diameters of the inhibition zones were measured, and isolates were categorized as susceptible, intermediate, or resistant based on the criteria established by the Clinical Laboratory Standards Institute (CLSI, 2016) and Charteris et al. (1998). L. plantarum LMG2003, L. sakei NCDO2714, B. longum, S. aureus ATCC 25923, and E. coli LMG3083 (ETEC) were included in the testing as quality control strains.

Isolates exhibiting resistance to three or more antibiotic classes were considered multidrug-resistant (MDR). The MAR index was also calculated using the formula described by Krumperman (1983): MAR index = number of antibiotics to which an isolate is resistant/total number of antibiotics tested.

Statistical analysis

Statistical analyses were conducted using SPSS version 26. Analysis of variance (ANOVA) was employed to evaluate differences between antibiotic groups. A p-value less than 0.05 was considered statistically significant.

Results

Enumeration and identification of Lactobacillus and Bifidobacterium species

A total of 10 probiotic food supplements including four sachets, three capsules, two tablets, and one drop are reported in Table 1 in terms of enumeration and identification of the contained lactobacilli and bifidobacteria. The agents and their labeled per dose composition were as follows: A1, Lactobacillus acidophilus, Lactobacillus rhamnosus (new name Lacticaseibacillus rhamnasus), Lactobacillus casei (new name Lacticaseibacillus casei), Bifidobacterium bifidum, 5×10⁹ CFU/dose; B2, Streptococcus thermophilus, B.bifidum, Bifidobacterium infantis, B. longum, Lactobacillus helveticus, L. acidophilus, Lactobacillus bulgaricus (new name Lactobacillus delbrueckii subsp. bulgaricus) (800 mg); C3, L. plantarum 299v, 10¹⁰ CFU/dose; C4, L. plantarum HEAL9, Lactobacillus paracasei (new name Lacticaseibacillus paracasei) n8700:2, 10⁹ CFU/dose; D5, L. acidophilus L1 , L. rhamnosus liobif , B. longum LBL-01 , Saccharomyces boulardii, 10×10⁹ CFU/dose; E6, S. thermophilus, L. plantarum, B. longum, Bifidobacterium breve, 2.3×10⁹ CFU/dose; E7, B. infantis, B.breve, B. longum, B. bifidum, 5×10⁸ CFU/dose; E8, Enterococcus faecium, L. acidophilus, L. rhamnosus, B. longum, B. bifidum, 2.5×10⁹; F9, Bifidobacterium animalis spp. lactis B94, 5×10⁹ CFU/dose; G10, L. acidophilus La-14, L. helveticus, L. rhamnasus Lr-32, B. longum BI-05, B. infantis Bi-26, Bifidobacterium lactis BI-04, Bacillus subtilis, B.bifidum Bb-06, L. plantarum Lp-115, L. bulgaricus Lb-87, Lactobacillus reuteri (new name Limosilactobacillus reuteri) 1E1, L. paracasei Lpc-37, Lactobacillus brevis Lbr-35, S. thermophilus St-21, 10×10⁹ CFU/dose. According to the label, 2 of 10 products contained monoculture bifidobacteria (E7, F9), 2 products contained monoculture lactobacilli (C3, C4), and 6 of them included mixed cultures.

The labeled number of cells and the counts (CFU) obtained for a unit dose (one capsule, one sachet, one tablet, or six drops) of each product can also seen in Table 1. Bacterial enumeration revealed that A1, E6, E7, and F9 have a lower content of viable cells than claimed. C3, C4, D5, E8, and G10 products were in agreement with their labels in the aspect of viable amount of bacteria. B2 packaging claimed a weight rather (800 mg) than a unit concentration and was found to have 6×10⁷ CFU/dose Lactobacillus spp., 1.2×10⁷ CFU/dose Bifidobacterium spp. None of the products that claimed to contain monocultures Bifidobacterium (E7, F9) were detected in bifidobacteria, but all products that claimed to contain monocultures Lactobacillus (C3, C4) contained the claimed culture concentration.

Thirteen presumptive lactobacilli or bifidobacteria strains were purely isolated and were phenotypically characterized. All the isolates were found to be Gram positive and catalase negative. Next, eight lactobacilli isolates and five bifidobacteria isolates were identified at the species level (Table 2). A1 and D5 evaluated were incompatible with the bacterial species claimed on the label. A1 product contained Lacticaseibacillus paracasei subsp. tolerans rather than L. acidophilus, L. rhamnasus, or L. casei mentioned on the label; similarly, the D5 contained B. infantis instead of B. longum stated on the label.

Lactiplantibacillus plantarum and Lacticaseibacillus rhamnosus were formerly known as Lactobacillus plantarum and Lactobacillus rhamnosus, respectively (Echegaray et al., 2022; Mathipa-Mdakane and Thantsha, 2022; Zheng et al., 2020). Table 2 shows that up-to-date names are not reflected in C3, C4, D5, E6, E8, and G10 products’ label information.

Antibiotic resistance

The antibiotic resistance of eight Lactobacillus and five Bifidobacterium probiotic strains from probiotic food supplements were analyzed by the disc diffusion method and were classified as either resistant (R), intermediate (I), or susceptible (S) based on the inhibition zone measured. A summary of the antibiotic resistance of Lactobacillus and Bifidobacterium isolates were reported in Table 3 and Table 4, respectively.

According to our results, all of the Lactobacillus strains were resistant to methicillin, cefoxitin, and vancomycin. Furthermore, resistance to cefazolin (37.5%), enrofloxacin (25%), ciprofloxacin (50%), norfloxacin (50%), kanamycin (37.5%), and trimethoprim (37.5%) was also observed, although at slightly lower levels. In contrast, none of the lactobacilli strains were resistant to ampicillin, chloramphenicol, clindamycin, erythromycin, gentamicin, penicillin G, rifampin, streptomycin, and tetracycline. L. paracasei subsp. tolerans strain was found to be resistant to cefoxitin, methicillin, and vancomycin, while being susceptible to ampicillin, cefazolin, chloramphenicol, ciprofloxacin, clindamycin, enrofloxacin, erythromycin, gentamicin, penicillin G, streptomycin, rifampin, tetracycline, and trimethoprim. L. acidophilus strain was resistant to cefoxitin, methicillin, trimethoprim, and vancomycin, while susceptible to ampicillin, cefazolin, chloramphenicol, ciprofloxacin, clindamycin, erythromycin, gentamicin, penicillin G, rifampin, streptomycin, and tetracycline. All L. plantarum strains were resistant to vancomycin, methicillin, cefoxitin, ciprofloxacin, and norfloxacin. Furthermore, the resistance to cefazolin (50%), enrofloxacin (50%), kanamycin (50%), and trimethoprim (25%) were also observed. However, none of the strains were found to be resistant to ampicillin, chloramphenicol, clindamycin, erythromycin, gentamicin, rifampin, penicillin G, tetracycline, and streptomycin. All L. rhamnosus strains were resistant to vancomycin, methicillin, and cefoxitin. In addition, the resistance to cefazolin (50%), kanamycin (50%), and trimethoprim (50%) were detected. Contrarily, none of the strains was found to be resistant to ampicillin, chloramphenicol, ciprofloxacin, clindamycin, enrofloxacin, erythromycin, gentamicin, norfloxacin, rifampin, penicillin G, tetracycline, and streptomycin.

All Bifidobacterium strains were resistant to methicillin and vancomycin. Frequent resistance was seen against cefazolin (80%), cefoxitin (80%), kanamycin (60%), and norfloxacin (60%). Furthermore, resistance to ampicillin (40%), clindamycin (20%), enrofloxacin (40%), trimethoprim (20%), and ciprofloxacin (20%) was also observed. None of the Bifidobacterium strains were found to be resistant against chloramphenicol, erythromycin, gentamicin, penicillin G, rifampin, streptomycin, and tetracycline.

B. bifidum strain was resistant to ampicillin, cefazolin, cefoxitin, clindamycin enrofloxacin, methicillin, norfloxacin, and vancomycin. B. longum strain was resistant to cefazolin, cefoxitin, kanamycin, methicillin, trimethoprim, and vancomycin. B. infantis strain was resistant to ampicillin, cefazolin, cefoxitin, kanamycin, methicillin, and vancomycin. B. breve strain was resistant to cefazolin, cefoxitin, ciprofloxacin, enrofloxacin, kanamycin, methicillin, norfloxacin, and vancomycin. B. lactis was resistant to methicillin, norfloxacin, and vancomycin.

The MDR defined as the resistance to three or more antimicrobial agents was found in all Lactobacillus and Bifdobacterium strains. L. paracasei subsp. tolerans strain was resistant to three antibiotics, one L. acidophilus and one L. rhamnosus strains were resistant to four antibiotics, one L. rhamnosus strain was resistant to five antibiotics, one L. plantarum strain was resistant to six antibiotics, and three L. plantarum strains were resistant to seven antibiotics. When the antibiotic resistance levels of the lactobacilli species were compared, L. plantarum strains showed a more resistant phenotype than other lactobacilli strains. Moreover, B.lactis strain was resistant to three antibiotics, B. infantis and B. longum strains were resistant to six antibiotics, and B. breve and B. bifidum strains were resistant to eight antibiotics. The majority of bifidobacteria isolates were resistant to six antibiotics and eight antimicrobials. In this study, MDR rates were found to be 100% in Bifidobacterium spp. and 100% in Lactobacillus spp. (Table 5). The data of the resistance in Bifidobacterium spp. and Lactobacillus spp. strains against antibiotics were statistically significant (p<0.001). The MAR index of the present study was notably unique for each isolate (Table 5). While 11 strains tested in the present study showed a MAR index of higher than 0.2, indicating a high-risk source of contamination, 2 strains showed a MAR index of lower than 0.2.

Discussion

Given the global surge in probiotic consumption, it is imperative to ensure that probiotic products are accurately labelled and contain well-documented strains, considering both safety and efficacy. The efficacy of probiotics often depends on strain-specific characteristics and the viability of the microorganisms upon reaching the gut (Korona-Glowniak et al., 2019; Temmerman et al., 2003). Manufacturers should adhere to rigorous labelling standards, identifying the genus, species, and strain of microorganisms in commercial probiotic formulations. Moreover, the labelled quantity of viable microorganisms should be guaranteed throughout the product’s shelf life under the specified storage conditions (FAO/WHO, 2002; Council for Responsible Nutrition and International Probiotics Association, 2017).

A significant discrepancy was observed between the labelled content and the actual microbial composition of five out of the ten probiotic products examined in this study. Our results corroborate previous studies by Aureli et al. (2010), Lugli et al. (2019), Kesavelu et al. (2020), and Syromyatnikov et al. (2022), which have reported inconsistencies between the labeled claims and the actual microbial content of probiotic products, including variations in the number, purity, type, and viability of microorganisms.

Our study’s findings align with those of cross-national investigations, which, as summarized below, have demonstrated discrepancies between the declared and actual microbial composition of commercial probiotic dietary supplements from various geographical regions. The study conducted by Temmerman et al. (2003) on 55 European probiotic products revealed that food supplements generally exhibited lower viable bacterial counts compared to dairy-based products. While a study by Drago et al. (2010) found that four out of thirteen probiotic supplements in the United States met their label claims, a subsequent study by Morovic et al. (2016) revealed that 33% of samples contained fewer viable bacteria than labelled or were mislabelled. A survey of probiotic dietary supplements in the United Kingdom conducted by Fredua-Agyeman et al. (2016) revealed that only three out of seven (43%) products contained the claimed concentration of probiotic bacteria. In addition, none of the multispecies products contained all of the labelled probiotic strains. Studies conducted in Bulgaria and China have demonstrated significant discrepancies between probiotic dietary supplements’ labels and actual microbial content. Respectively, Marinova et al. (2019) found that only 10 out of a certain number of products contained the claimed 10⁸ to 10¹⁰ CFU/g of probiotic bacteria, and none of the commercial products contained all labelled LAB species. In addition, some products were found to contain unacceptable microorganisms. Similarly, Ullah et al. (2019) reported that 29.41% of the capsule and sachet-based probiotic products in their study contained inaccurate or lower CFU counts, while 23.52% did not comply with the labelled microbial composition. Recent research by Anisimova et al. (2022) revealed inconsistencies between certain labelled probiotic products and the actual microbial composition. Some samples were labelled as multispecies formulations but were found to contain only a single species of lactobacilli. In addition, other samples contained species that differed from those claimed by the manufacturers. Similarly, a study conducted by Gundogdu et al. (2023) on probiotic products in Türkiye found discrepancies between the label and actual content in 17 examined products. These findings align with our results, highlighting the prevalence of mislabelling and misidentification of probiotic strains in the market.

Our findings align with previous studies by Chen et al. (2014) and Lewis et al. (2016), which demonstrated that Bifidobacterium species were either absent or present in low numbers in products labeled as containing these bacteria. Similarly, Aureli et al. (2010) reported the frequent mislabeling of probiotic products, with B. bifidum often claimed but rarely detected, and when detected, found to be nonviable.

Suboptimal processing techniques can compromise the viability of microorganisms, while inadequate packaging, storage, and transportation conditions can further reduce bacterial survival. Consequently, careful consideration must be given to selecting strains that are robust to manufacturing processes or optimize manufacturing processes to accommodate sensitive strains. Bifidobacterial species, in particular, are more susceptible to adverse manufacturing and storage conditions than other bacterial strains (Drago et al., 2010; Fredua-Agyeman et al., 2016).

Probiotic bacteria can exhibit either resistance or sensitivity to antibiotics, depending on the presence or absence of resistance genes in their genome, plasmid-based antibiotic resistance genes, or intrinsic resistance mechanisms. The transfer of antibiotic resistance determinants from probiotic bacteria to the intestinal microbiota and potential opportunistic pathogens poses a significant health concern. Consequently, the long-term consumption of probiotic dietary supplements and foods warrants careful consideration because of the potential risk of disseminating antibiotic resistance (Jose et al., 2015; Shahali et al., 2023).

In the present study, the antibiotic resistance profiles of eight Lactobacillus strains and five Bifidobacterium strains isolated from probiotic food dietary supplements were evaluated. Notably, all tested strains demonstrated resistance to multiple antibiotics (three or more), revealing a pattern of MDR. These findings are consistent with previous research by Anisimova et al. (2022) and Sharma et al. (2016), which also reported widespread antibiotic resistance among probiotic lactobacilli strains.

All Lactobacillus strains in our study exhibited resistance to vancomycin, consistent with the findings of Wong et al. (2015), Wang et al. (2022b), and Zavišić et al. (2023). However, in contrast to our results, lower levels of resistance to vancomycin were reported by Sharma et al. (2016) and Anisimova et al. (2022), who observed resistance rates of 76% and 79%, respectively. In alignment with our findings, Han et al. (2015) determined that all L. plantarum and L. rhamnosus strains exhibited resistance to vancomycin. Similarly, Hammad and Shimamoto (2010) reported high levels of vancomycin resistance in L. plantarum strains. The resistance of lactobacilli to vancomycin is widely regarded as intrinsic, chromosomally encoded, and nontransferable (Anisimova and Yarullina, 2019). Zaviši´c et al. (2023) reported that all Lactobacillus isolates were resistant to cefoxitin, consistent with the findings of our study. However, in contrast to our results, Sharma et al. (2016) observed lower levels of resistance to methicillin. Contrary to the findings of Zaviši´c et al. (2023), none of the Lactobacillus strains isolated in our study exhibited resistance to penicillin. Similarly, Sharma et al. (2016) reported that most Lactobacillus strains displayed low levels of resistance to penicillin. In agreement with our results, Guo et al. (2017) found that all Lactobacillus strains were susceptible to gentamicin, whereas Turhan and Enginkaya (2016) reported that 20% of Lactobacillus spp. isolates were resistant to this antibiotic. Furthermore, the susceptibility of Lactobacillus strains to erythromycin and clindamycin observed in our study aligns with the findings of Sharma et al. (2016) and Guo et al. (2017). Turhan and Enginkaya (2016) also found that all Lactobacillus isolates were susceptible to chloramphenicol and erythromycin, consistent with our results. However, they reported higher resistance rates to tetracycline (20%), ampicillin (20%), gentamicin (20%), and ciprofloxacin (80%) compared to our findings. The resistance of Lactobacillus isolates to cefazolin in our study was 37.5%, closely aligning with the 32% reported by Anisimova et al. (2022). In contrast, Han et al. (2015) reported higher incidences of resistance to penicillin G (37.9%), ampicillin (34.5%), streptomycin (93.1%), kanamycin (100%), tetracycline (10.3%), chloramphenicol (10.3%), gentamicin (86.2%), and rifampin (34.5%). In our study, L. acidophilus strains were susceptible to ampicillin, chloramphenicol, erythromycin, rifampin, and tetracycline, consistent with the findings of Zhou et al. (2005) and Turhan and Enginkaya (2016). Similarly, Zhou et al. (2005) and Temmerman et al. (2003) observed that L. acidophilus strains were susceptible to penicillin, aligning with our findings. However, Zhou et al. (2005) reported resistance to kanamycin and streptomycin in L. acidophilus strains, which contradicts our results. Conversely, Selvin et al. (2020) found L. acidophilus to be resistant to ampicillin and L. rhamnosus resistant to erythromycin, contrasting with our study. Similarly, Kim et al. (2020) observed resistance in L. rhamnosus MG316 to kanamycin and chloramphenicol, while in our study, all L. rhamnosus strains were sensitive to chloramphenicol, and only one of two strains exhibited resistance to kanamycin. Anisimova and Yarullina (2019) reported that none of the L. plantarum strains were resistant to ampicillin, erythromycin, chloramphenicol, or tetracycline and that all strains exhibited resistance to ciprofloxacin and norfloxacin, consistent with our results. In addition, 10 of 11 strains were resistant to vancomycin, aligning with our findings. Gupta and Tiwari (2014) observed that the L. plantarum LD1 strain was resistant to kanamycin but sensitive to tetracycline, erythromycin, chloramphenicol, and gentamicin, which aligns with our findings that all L. plantarum strains were sensitive to erythromycin, chloramphenicol, and gentamicin. However, in our study, two of four L. plantarum strains exhibited resistance to kanamycin.

All Bifidobacterium strains in this study exhibited resistance to vancomycin, consistent with the findings of Zuo et al. (2016). However, this contrasts with the results of Kim et al. (2018), who reported that all Bifidobacterium spp. strains were susceptible to vancomycin. D’Aimmo et al. (2007) observed high resistance levels to kanamycin in bifidobacteria, aligning with our finding that 60% of the tested Bifidobacterium strains were resistant to kanamycin. In our study, all Bifidobacterium strains were susceptible to chloramphenicol, erythromycin, gentamicin, rifampin, and tetracycline. This contrasts with the results of Xu et al. (2018), who reported resistance rates of 8.7% to chloramphenicol, 4.35% to erythromycin, 13.04% to gentamicin, 10.87% to rifampin, and 43.48% to tetracycline among Bifidobacterium strains. In agreement with our findings, Kim et al. (2018) reported that all Bifidobacterium strains were susceptible to chloramphenicol, rifampin, and erythromycin, while exhibiting general resistance to kanamycin. Conversely, Yasmin et al. (2020) found that all Bifidobacterium strains were resistant to gentamicin, kanamycin, and most were resistant to streptomycin and rifampin, though all were susceptible to tetracycline, consistent with our results. Similarly, Moubareck et al. (2005) demonstrated that none of the tested Bifidobacterium strains, regardless of species, exhibited resistance to penicillin G, in alignment with our findings. Zuo et al. (2016) reported universal susceptibility of isolated bifidobacteria to ampicillin, while our study found that 40% of Bifidobacterium strains were resistant to ampicillin. Turhan and Enginkaya (2016) observed that Bifidobacterium strains were resistant to vancomycin, tetracycline, ampicillin, and ciprofloxacin but sensitive to chloramphenicol, erythromycin, and gentamicin. In contrast, none of the Bifidobacterium strains in our study exhibited resistance to tetracycline. Moubareck et al. (2005) found that one of fourteen B. longum strains and one of six B. breve strains were resistant to cefoxitin, while in our study, all Bifidobacterium species except B. lactis exhibited resistance to cefoxitin. Dioso et al. (2020) reported resistance to erythromycin, kanamycin, and vancomycin in B. breve strains, findings consistent with ours except for erythromycin. Similarly, B. breve strains were resistant to ciprofloxacin in our study, aligning with the observations of Awasti et al. (2016). Kim et al. (2020) found that B. breve MG729 exhibited resistance to tetracycline, whereas Kim et al. (2018) observed high antibiotic resistance to gentamicin and tetracycline in B. bifidum BGN4 and B. longum BORI. However, in our study, none of the Bifidobacterium strains exhibited resistance to tetracycline or gentamicin. The susceptibility of B. bifidum strains to erythromycin, tetracycline, gentamicin, and penicillin G in our study aligns with the findings of Drago et al. (2013). In contrast, Wei et al. (2012) reported that B. longum JDM301 was intrinsically resistant to ciprofloxacin, gentamicin, and streptomycin while being susceptible to vancomycin and trimethoprim. In addition, Wei et al. (2012) observed susceptibility to chloramphenicol, erythromycin, ampicillin, and rifampin, consistent with our findings. All B. lactis strains were reported to be resistant to kanamycin, gentamicin, and streptomycin and susceptible to ampicillin and rifampin by Zhou et al. (2005), findings consistent with ours. Similarly, Temmerman et al. (2003) found that B. lactis strains were susceptible to tetracycline, chloramphenicol, erythromycin, and penicillin G, in agreement with our study. However, in our study, B. lactis strains exhibited resistance to vancomycin, with levels of resistance comparable to those reported by Temmerman et al. (2003) (50%) and Zhou et al. (2005) (33%).

In this study, MDR rates were found to be significantly high among the probiotic strains tested. To the best of our knowledge, this is the first study to report MDR rates in probiotic cultures isolated from commercial probiotic dietary supplements (Table 5). A key novelty of the present research lies in the calculation and reporting of the multiple antibiotic resistance (MAR) index for probiotic strains isolated from probiotic dietary supplements in Türkiye for the first time. The MAR index values in this study were distinctly unique for each isolate (Table 5). The MAR index is a crucial metric used to assess the level of antibiotic resistance in microorganisms, particularly in probiotic strains, which are commonly used in dietary supplements. This index provides an effective way to evaluate the extent of resistance, which can be a critical factor when selecting probiotics for use in commercial applications (Duche et al., 2023). Several studies have reported that MAR index greater than 0.2 indicates a high-risk antibiotic-exposed source (Ayandele et al., 2020; Korzeniewskan et al., 2013; Okeke et al., 2005). A study by Duche et al. (2023) showed that while probiotic strains exhibited some degree of resistance to antibiotics, their MAR index remained relatively low, suggesting a limited risk of contributing to overall antimicrobial resistance. However, a higher MAR index in probiotic strains may raise concerns about their safety and long-term use, especially when taken alongside antibiotics (Haryani et al., 2023). Notably, 11 of the tested strains exhibited a MAR index greater than 0.2, indicating a high-risk source of contamination, whereas 2 strains showed a MAR index lower than 0.2. The high MAR index that was reported in the present work thus indicated a high-risk potential associated with fermented food products that could threaten human health. These findings underscore the critical importance of routine and continuous monitoring of antibiotic resistance patterns in probiotic strains used in probiotic dietary supplements. Such measures are essential to ensure the safety and efficacy of probiotic formulations. Further research is warranted to thoroughly investigate the antibiotic resistance determinants and mechanisms in these isolates to confirm the safety of dairy-based probiotic formulations.

Conclusions

Results from this study, aimed at evaluating the compositional quality and antibiotic resistance profiles of commercial probiotic food supplements in Türkiye, revealed, through both conventional culture-based and molecular methods, that some products did not meet the label claims regarding the quantity and variety of viable microorganisms. Results from the second part of the study, which investigated the phenotypic antibiotic resistance profiles of Lactobacillus spp. and Bifidobacterium spp. strains isolated from probiotic supplements, demonstrated a high prevalence of antibiotic resistance. Despite rigorous selection criteria for candidate microorganisms based on their safety, and technological suitability, the findings indicate significant concerns in products reaching consumers. In light of the growing popularity of probiotic dietary supplements, particularly during the COVID-19 pandemic, there is an urgent need to re-evaluate the regulations governing these products to ensure the safety and efficacy of the probiotic microorganisms they contain. The findings of this study underscore the critical need for enhanced quality control measures within the probiotic industry. To address the identified shortcomings and ensure the safety and efficacy of probiotic products, the following recommendations are proposed:

1. A comprehensive review of manufacturing processes is necessary to identify and implement strategies that minimize microbial losses, particularly for sensitive strains such as bifidobacterium, throughout the product shelf life.

2. Robust regulatory frameworks should be established to mandate the monitoring of the quantity and diversity of probiotic microorganisms in supplement formulations and the antibiotic resistance profiles of the included species.

3. Product labels for probiotic dietary supplements should clearly communicate the potential health benefits associated with the included microbial strains, emphasizing strain-specific effects supported by scientific evidence.