The plant material used in this study was the bark of Aesculus hippocastanum. The plant was collected in June 2021 near the Medical Institute of the Peoples’ Friendship University of Russia (MG23 + 9H Obroutchevski, Moscow, Russia). After harvest, the plant was taken directly to the laboratory where it was dried at 37 ${}^{\circ }$C until the weight was constant. The dry matter content was calculated; the plant was grinded and the powders with particle sizes lower than 1 mm were stored in a sterile airtight container until further use.

The microorganisms used for the screening of antimicrobial activity consisted of five Gram positive bacteria and 5 Gram negative. The five Gram + were Kocuria rhizophila 1542, Enterococcus avium 1669, Staphylococcus simulans 5882, Corynebacterium spp 1638 and Enterococcus faecalis 5960 while the five Gram—were Proteus mirabilis 1543, Morganella morganii 543, Citrobacter freundi 426, Acynetobacter baumannii 5841 and Achromobacter xylosoxidans 4892. Staphylococcus aureus ATCC 6538 and Escherichia coli ATCC 25922 were used as standard Gram + and Gram - respectively. All strains were provided by the Department of Microbiology and Virology of the Peoples’ Friendship University of Russia.

Dimethyl sulfoxide (DMSO) was purchased from BDH Laboratories, VWR International Ltd., USA. We also used BHIB (Brain Heart Infusion Broth) (HiMedia™ Laboratories Pvt. Ltd., India), Muller Hinton Agar (MHA HiMedia™ Laboratories Pvt. Ltd., India), Sabouraud Dextrose Broth (SDB, HiMedia™ Laboratories Pvt. Ltd., India) and all other reagents and chemicals used were of analytical grade.

Ethanolic solution (80%, v/v) and distilled water were used as solvents because they have been reported to be efficient solvents for the extraction of bioactive compounds in the medicinal plants used in this study. As we described in our previous investigation [18] thirty grams (30 g) of vegetal material was weighed and added to 270 mL of the solvent in separate conical flasks. The flasks were covered tightly and were shaken at 200 rpm for 24 h and 25 ${}^{\circ }$C in a shaker incubator (Heidolph Inkubator 1000 coupled with Heidolph Unimax 1010, Germany). The mixtures were then filtered by vacuum filtration, using Whatman filter paper № 1 then concentrated at 40 ${}^{\circ }$C in rotary evaporator (IKA RV8) equipped with a water bath IKA HB10 (IKA Werke, Staufen, Germany) and a vacuum pumping unit IKA MVP10 (IKA Werke, Staufen, Germany). To avoid losses, the extracts were collected when the volumes were small enough and placed in petri dishes previously weighed and then incubated open at 40 ${}^{\circ }$C until complete evaporation. The final dried crude extracts were weighed. Extraction volume and mass yield were determined using the following formulas:

(1)$$\text{Volume yield}(\%)=\frac{\text{Volume of the extract after filtration}(\mathrm{ml})}{\text{Initial solvent volume}(\mathrm{ml})}\times 100$$

(2)$$\text{Mass yield}(\%)=\frac{\text{Mass of extracted plant residues}(\mathrm{g})}{\text{Mass of plant raw sample}(\mathrm{g})}\times 100$$

For each plant extract, the crude extract was dissolved in the required volume of DMSO (5%, v/v) to achieve a concentration of 128 mg/mL. The extracts were sterilized by microfiltration (0.22 $\mu$m) and the solution obtained was used to prepare the different concentrations used in the analytical process.

Bacteria were cultured for 24 h at 37 ${}^{\circ }$C in 10 mL of BHI broth. After incubation, the cells were collected by centrifugation (7000 g, 4 ${}^{\circ }$C, 10 min), washed twice with sterile saline, resuspended in 5 mL of sterile saline to achieve a concentration equivalent to McFarland 0.5 using DEN‑1 McFarland Densitometer (Grant‑bio).

The modified Kirby–Bauer’s disk method described in our previous study [22] was used to study the antibiotic sensitivity of the bacterial strains, and the following eight antibiotics disks were used: amoxicillin, 30 $\mu$g/disk; ampicillin, 25 $\mu$g/disk; cefazolin, 30 $\mu$g/disk; cefazolin/clavulanic acid, 30/10 $\mu$g/disk; 30 $\mu$g/disk; ceftriaxone, 30 $\mu$g/disk; ciprofloxacin, 30 $\mu$g/disk; Fosfomycin, 200 $\mu$g/disk; imipenem (IMP), 10 $\mu$g/disk; nitrofurantoin, 200 $\mu$g/disk; tetracyclin (TE), 30 $\mu$g/disk and trimethoprim, 30 $\mu$g/disk. The inhibition diameters were measured and interpreted then referred to the Clinical & Laboratory Standards Institute [23]. Resistance R, Intermediate I, and Sensitive S interpretations were obtained automatically using algorithms written in Excel software [Microsoft Office 2016 MSO version 16.0.13628.20128 (32 bits), USA] with the parameters described in Table 1 [10].

The dry matter, water content, ethanolic (EE) and distilled water extract (AE) yields of Aesculus hippocastanum bark are presented in Table 2. We found that A. hippocastanum bark possessed a dry matter content of 65.73%. Dry matter is an indicator of the amount of constituents (excluding water) in the sample tested. The drying allowed the removal of water from the plant in order to standardize the extraction and to make this work reproducible if authors from other regions project to work with the same plant. Furthermore, we obtained an extraction volume yield of 74.07% with distilled water as the solvent and 77.77% with ethanol. The difference in extraction volume yields can be explained by losses during the extraction process. Indeed, as reported by [18] we noticed that the filtration of ethanolic extracts was faster (less than 5 minutes for 300 mL) compared to the aqueous extract which took much longer (more than 50 minutes on average for 300 mL) and required in average 6 filter changes. Despite the filtration, the EAs still looked cloudy while the EEs were completely clear. This residue cloudiness could therefore explain the higher mass yield in AE (24.3%) compared to EE (13.4%). Several authors [17, 18, 26, 27] reported observations similar to our findings while others [28, 29] found opposite results. Therefore, as Arsene et al. [18] pointed out in their previous work, the extraction performance depends on several factors, including the method used, extraction time, the solvents, and the equipment used. In addition, Mouafo et al. [17] reported that the high yields of phytoconstituent does not necessarily imply better antimicrobial activity and this was further assessed in the present work by evaluating the antimicrobial activity of each of the extracts using the well diffusion method and the determination of minimum inhibitory concentrations (MIC) and minimum bactericidal concentrations (MBC).

The susceptibility of uropathogenic tests to eleven (11) antibiotics was evaluated by the Kirby Bauer disc method, the Multidrug resistance Index (MDR) was calculated, and the results were reported in Table 3. As shown in Table 3, no bacteria were resistant to amoxiclav, imipenem and ceftriaxone while 7/12 were resistant to ampicillin, 6/12 to trimethoprim, 5/12 to tetracycline and ceftazidime/clavulanic acid, 4/12 to ceftazidime, 3/12 to fosfomycin, 2/12 to nitrofurantoin and 1/12 to ciprofloxacin. Interestingly, both standard bacteria (E. coli ATCC 25922 and S. aureus ATCC 6538) were sensitive to all antibiotics while the clinical strains were resistant to at least one antibiotic. Among clinical strains, MDRs ranged from 0.09 to 0.45. K. rizophilia 1542 and Corynebacterium spp 1638 were the most resistant bacteria with MDRs of 0.45 each. The results observed with these clinical strains are similar to those reported in other studies [10, 11]. While the strongest resistance was observed on ampicillin (which is an antibiotic from the $\beta$-lactams family, from the group A of the penicillins), no resistance was observed in amoxiclav which is a derivative of amoxicillin (antibiotic of the beta-lactam family, of the aminopenicillin group) of the same family, with the only difference that the latter is combined with clavulanic acid ($\beta$-lactamase inhibitor). This shows that antibiotic adjuvants can also play an important role in combating resistant germs [30]. Furthermore, similarly to our findings, other studies have reported low resistance of uropathogens to imipenem and ceftriaxone [11, 31]. In addition, the sensitivity of all strains to amoxiclav, imipenem and ceftriaxone demonstrates that these 3 antibiotics are effective on a wide range of bacteria, including clinical strains with high resistance to other antibiotics. Therefore, these antibiotics which have kept a good activity against various microorganisms should be carefully used and administered only under prescription and after a prior antibiogram.

Fig. 2 presents the inhibition diameters of ethanolic (EE) and aqueous (AE) extract of Aesculus hippocastanum bark on the tested microorganisms. Except AE on Proteus Mirabilis 1543 and Enterococcus faecalis 5960 (0 mm), both AE and EE were active on all microorganisms tested with inhibition diameters (mm) which ranged from 5.5–10.0 for AE and 8.0–14.5 for EE.

Fig. 2.

Inhibition diameter of ethanolic and aqueous Aesculus hippocastanum bark extract on tested uropathogenic bacteria.

The ethanolic extracts (EE) were overall more active than the aqueous ones. Consequently, this means that ethanol extracts more compounds with antimicrobial properties compared to water although we found above that the extraction yields with water was higher than that with ethanol. Several authors have reported that compounds with antimicrobial activity such as flavonoids, polyphenols, tannins and alkaloids are generally insoluble in water but soluble in ethanol [32, 33]. Other authors such as Arsene et al. [18], Mouafo et al. [17] and Evbuomwan et al. [34] also pointed out that ethanol extracts more antimicrobial compounds from plant materials as opposed to water. In addition, we found that extracts of A. hippocastanum bark were both active against Gram + bacteria as against Gram - bacteria. To our knowledge, the antibacterial properties of this plant have not yet been investigated but our findings suggest that A. hippocastanum bark has constituents exhibiting a broad-spectrum antimicrobial.

Most of the research conducted on A. hippocastanum focused on its properties to regulate circulatory system imbalances and relieve attacks of hemorrhoids [19]. In a study conducted by Owczarek et al. [19] on the composition of A. hippocastanum bark, it has been reported that this plant presents mainly two groups of compounds: coumarins and proanthocyanidins. Among the coumarins there was mainly esculin (over 17%) and fraxin (over 7%) while among the proanthocyanidins, the authors found epicatechin (over 6%) and procyanidin A2 (over 5%) (Fig. 1) [19]. Owczarek et al. [19] finally concluded that, in total, about 40% of the A. hippocastanum bark extract could be attributed to simple phenolics compounds detectable by LC-PDA. Unfortunately, due to limited ressources we were unable to assess the composition of our extracts in order to compare with the data in the literature. However, we can hypothesize that the antimicrobial properties of A. hippocastanum bark can be attributed to all its components or specifically to esculin [35], to fraxin [36] or proanthocyanidins [37] because these compounds (from other plants) have been reported to have antimicrobial properties. Notwithstanding our findings, further investigations are required to confirm or refute our hypothesis.

After evaluating the antibacterial properties of aqueous (AE) and ethanolic (EE) extracts of Aesculus hippocastanum bark against the uropathogens tested using the well diffusion, we investigated the minimal inhibitory concentrations (MIC) and minimal bactericidal concentrations (MBC) of the two extracts. MIC and MBC are two very important elements in the search for new antimicrobials and respectively provide the minimum concentrations required to inhibit or kill the microorganism tested. Table 4 presents the MIC, the MBC, and the MBC/MIC ratio of our 2 extracts on the uropathogens investigated. Similarly to the inhibition diameters, ethanolic extracts (EE) showed the best antimicrobial activity with the lowest MIC and MBC. The MICs of EEs varied from 1–4 mg/mL while those of EAs varied from 4–6 mg/mL. Almost all the MBCs of AEs were indeterminate ($>$64 mg/mL) while those of EE were successfully determined.

With water as a solvent, the highest antimicrobial activity was observed against E. faecalis 5960, S. aureus ATCC 6538 and C. freundi 426. Although the MIC and MBC of AE were high against these bacteria, AE was found to be bactericidal against S. aureus ATCC 6538 and C. freundi 426 since the MBC/MIC ratio was 4 for both bacteria. Indeed, Mondal et al. [25] reported that when the ratio MBC/ MIC is $\ge$16, the antibacterial efficacy of the test agent is considered as bacteriostatic, whereas MBC/MIC $\le$4 indicates bactericidal activity. Similarly, although the MICs of EEs were relatively low, the MBCs of this extract were quite high, and we concluded that the antibacterial activity was overall bacteriostatic (MBC/ MIC $\ge$16) against most bacteria except against some Gram-positive bacteria such as S. simulans 5882, Corynebacterium spp 1638 and S. aureus ATCC 6538. This difference in the activity (although not significant) of EE between Gram positive and Gram negative can be ascribed to the cell wall structure of Gram - bacteria, which differs from the structure of Gram + bacteria with a thin layer of peptidoglycan, the presence of a periplasm, and ease of exchange on the plasma membrane [18]. Furthermore, according to the classification of Kuete [38], the different extracts could be considered as deserving a weak antimicrobial activity independently of the extraction solvent and the tested strain as they scored a MIC value higher than 0.625 mg/mL. Therefore, further studies are needed to evaluate the antibacterial activity of A. hippocastanum bark extracts with other solvents and to assess their synergy with other antibiotics, as the weak antimicrobial activity observed here does not allow us to recommend our plant extracts as such for a possible use in the fight against antibiotic resistance and the management of urinary tract infections.