Do South African medicinal plants used traditionally to treat infections respond differently to resistant microbial strains?

Do South African medicinal plants used traditionally to treat infections respond differently to resistant microbial strains?

South African Journal of Botany 112 (2017) 186–192 Contents lists available at ScienceDirect South African Journal of Botany journal homepage: www.e...

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South African Journal of Botany 112 (2017) 186–192

Contents lists available at ScienceDirect

South African Journal of Botany journal homepage: www.elsevier.com/locate/sajb

Do South African medicinal plants used traditionally to treat infections respond differently to resistant microbial strains? S. van Vuuren ⁎, T. Muhlarhi Department of Pharmacy and Pharmacology, Faculty of Health Sciences, University of the Witwatersrand, 7 York Road, Parktown, 2193, South Africa

a r t i c l e

i n f o

Article history: Received 18 April 2017 Received in revised form 10 May 2017 Accepted 22 May 2017 Available online xxxx Edited by B Ncube Keywords: Aqueous extract Antibiotic Bacteria Essential oil Gram-negative Gram-positive Organic extract Resistance

a b s t r a c t Currently antimicrobial resistance is increasing at an alarming rate. Exposure to resistant strains hinders treatment outcomes both in rural and hospital settings. Thus, the aim of this study was to investigate five frequently used South African medicinal plants (Artemisia afra, Lippia javanica, Osmitopsis asteriscoides, Croton gratissimus and Tetradenia riparia) and test these against resistant bacterial strains (Enterococcus faecalis, Staphylococcus aureus, Bacillus cereus, Klebsiella pneumoniae, Acinetobacter baumannii, Pseudomonas aeruginosa, Escherichia coli and Serratia marcescens) and comparatively evaluate efficacy with a reference strain. The conventional antibiotic ciprofloxacin was used as a positive control to also compare susceptibility of the various strains. Most plant samples demonstrated similar or better activity against the resistant strains. A general trend demonstrated that the organic extracts followed by the essential oils were able to withstand resistant strains better than the antibiotics which showed reduced susceptibility. This demonstrates great promise as natural products provide an alternative to fighting the onslaught of antibiotic resistance. © 2017 SAAB. Published by Elsevier B.V. All rights reserved.

1. Introduction Currently antimicrobial resistance is a global crisis, increasing at an alarming rate. The last few decades have shown an increase in clinical and community acquired multidrug resistant infections which have had an impact on treatment regimens (Canton and Morosini, 2011). It is estimated that by the year 2050, infections by antimicrobial resistant organisms will be the leading cause of death worldwide (O'Neill, 2014). Important bacterial strains conveying resistance include Staphylococcus aureus, Pseudomonas aeruginosa, Acinetobacter baumannii and Klebsiella pneumoniae amongst others (Warnke et al., 2009; Khan and Zarrilli, 2012; Potron et al., 2015). In South Africa, antimicrobial resistance has reached alarming proportions. Some statistical impact factors include studies that show more than 50% of all hospital-acquired S. aureus infections were of methicillin resistance origin (Bamford et al., 2011). In an editorial entitled “Wake up, South Africa! The antibiotic horse has bolted” (Mendelson et al., 2012), the very title leaves no subtlety as to the huge problem South Africa is facing with respect to antimicrobial resistance. To address the crisis and challenges posed by the growing global resistance of micro-organisms to conventional antimicrobials, research ⁎ Corresponding author. E-mail address: [email protected] (S. van Vuuren).

http://dx.doi.org/10.1016/j.sajb.2017.05.027 0254-6299/© 2017 SAAB. Published by Elsevier B.V. All rights reserved.

has adapted a growing interest in examining alternatives such as natural products (Al-Mariri and Safi, 2014; Harvey et al., 2015; Moloney, 2016). Some recent studies to examine natural products and resistant strains include the investigation of Uapaca togoensis and related compounds when tested against Gram-negative multi-drug resistant phenotypes (Seukep et al., 2016). Eucalyptus camaldulensis has been studied against multi-drug resistant Acinetobacter baumannii (Knezevic et al., 2016). Antibacterial effects of cinnamon oil have been investigated against carbapenem resistant nosocomial Acinetobacter baumannii and Pseudomonas aeruginosa isolates (Kaskatepe et al., 2016). The antimicrobial activity of Artemisia judaica (essential oil) against clinical multi-drug resistant bacteria (Benmansour et al., 2016) has been examined and studies on crude extracts against multiple resistant urinary tract infections (Mishra et al., 2017) have also had some attention. These are just a few studies that have focused on resistant strains. South Africa is home to more than 3000 species of medicinal plants (Van Wyk et al., 1997), and many of these have shown great antimicrobial potential (Van Vuuren, 2008; Van Vuuren and Holl, submitted for publication). In spite of the number of publications dedicated to natural products and resistant strains, very little attention has been given to resistant strains and medicinal plants from a South African perspective. In a very recent review of antimicrobial studies and South African natural products (Van Vuuren and Holl, submitted for publication), only a few papers (Lall and Meyer, 1999; Heyman et al., 2009; Bisi-Johnson et al.,

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2011; Njume et al., 2011a, 2011b; Nielsen et al., 2012; Mabona et al., 2013; Khan et al., 2014; Nciki et al., 2016) included resistant strains and none of these studies comparatively evaluated the antimicrobial effects of natural products with conventional antibiotics. It is well known that medicinal plants are not as potent as single compound antibiotic drugs. Medicinal plants are considered active when efficacy (minimum inhibitory concentrations [MIC]) under 100 μg/mL are evident (Van Vuuren and Holl, submitted for publication). The effective MIC values for antibiotics varies between strains and have recommended acceptable breakpoint values (Andrews, 2004; CLSI guidelines, 2012), which are clearly much lower and in microgram quantities. Even though there is a large variation in potency, medicinal plants are more readily available, cheaper and cultural reasons validate selection and choice. Even though the efficacy of medicinal plants do not show the potency that single compound antibiotics offer they may be able to demonstrate more resilience against resistant microbial strains when compared to conventional antibiotics. This may in the future become a vital advantage in treating infections that harbour resistant strains. With this in mind, the overall aim of this study was to compare the antimicrobial responses of resistant strains with that against a reference strain when exposed to indigenous South African medicinal plants. Furthermore the comparative response observed when exposed to conventional antibiotics was explored. 2. Methodology 2.1. Plant selection and preparation A selection of five common South African medicinal plants [Artemisia afra Jacq. ex Willd. (Asteraceae), Osmitopsis asteriscoides (L.) Less. (Asteraceae), Lippia javanica Spreng. (Verbenaceae), Croton gratissimus Burch. (Euphorbiaceae) and Tetradenia riparia (Hochst.) Codd (Lamiaceae)] was chosen based on popularity, traditional use to treat infections and the fact that they are aromatic which allowed for the inclusion of the essential oils in the study. These plants are all antimicrobially well studied (Mangena and Muyima, 1999; Viljoen et al., 2003; Viljoen et al., 2005; Van Vuuren and Viljoen, 2008; Shikanga et al., 2010; Suliman et al., 2010; York et al., 2012), yet little attention has been given to how these medicinal plants respond to resistant bacterial strains. Plant species (with the exception of O. asteriscoides which was collected from a population near Betty's Bay in the South Western Cape region of South Africa) were collected from the Walter Sisulu botanical gardens with permission and assistance from Mr. Andrew Hankey, Specialist horticulturist and plant conservationist. Identification was confirmed and voucher specimens prepared and stored in the Department of Pharmacy and Pharmacology, University of the Witwatersrand. Dried plant samples were ground and immersed in a 1:1 mixture of dichloromethane and methanol. This organic extract was selected for the ability to extract a combination of both polar and nonpolar compounds. The plant samples in solvent was retained in the shaker incubator (Labcon) at 37 °C for 24 h. Aqueous extracts were prepared by submerging the macerated plant material in sterile distilled water which was kept at ambient temperature overnight. Thereafter the extracts were filtered, stored at −80 °C and lyophilized (VirtTis) (Van Vuuren and Viljoen, 2006). For the essential oils, a known quantity of weighed fresh leaf material was subjected to hydrodistillation using a Clevenger-type apparatus. After 3 h, the essential oil was collected, weighed and stored in amber bottles at 4 °C. 2.2. Culture strains Bacterial strains (Table 1) included in the study were selected on the basis of emerging antimicrobial resistance patterns. These included the “ESKAPE” pathogens which have come to encompass Staphylococcus aureus, Klebsiella pneumoniae, Pseudomonas aeruginosa, Enterococcus spp. and Acinetobacter spp. The selected strains represent pathogens

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with growing multidrug resistant virulence on a global scale. The number of test strains included was based on availability. Where clinical resistance is common, more strains were included. These included Enterococcus faecalis (five resistant strains), Staphylococcus aureus (eight resistant strains) and Klebsiella pneumoniae (three resistant strains). A non-resistant reference strain was included for each strain type to compare the antimicrobial response. Resistant strains were obtained from various sources as indicated in b–d (Table 1). All cultures were confirmed pure before commencement of the study and resistance patterns were predetermined by the relevant suppliers. Eight different bacterial species (three Gram-positive and five Gram-negative strains) comprising of a total of 32 different strains were included in the study (Table 1). Culture purity was monitored throughout the study by using the streak plate method. Sterility of media was confirmed by incubating un-inoculated broth with the tests. 2.3. The minimum inhibitory concentration (MIC) method The MIC method was modified from Eloff (1998) and is in accordance with the CLSI guidelines (2012). All plant samples (essential oils, organic and aqueous extracts) were prepared to a starting concentration of 32 mg/mL. The organic extracts and essential oils were diluted in acetone. The aqueous plant samples were diluted in sterile water. The positive control was prepared by diluting ciprofloxacin to a concentration of 0.01 mg/mL in sterile water. This was an important comparator to include as a response of inhibitor (either tests or antibiotic) was crucial to determine if resistant strains respond differently. A negative control was included to determine if solvent (acetone) exhibited any antimicrobial effects. The method, in brief, involved incorporating 100 μL of test sample and or control together with 100 μL of sterile Tryptone Soya broth (TSB) into the top row of a micro-titre plate. The test samples were then serially diluted in TSB to give concentrations of 8, 4, 2, 1, 0.5, 0.25, 0.125 and 0.075 mg/mL. A McFarland standard (approximately 1 × 106 colony forming units [CFU]/mL) was used as a guide to prepare cultures and 100 μL added into all the wells of each plate. Each microtitre plate was sealed with a sterile adhesive film to prevent evaporative loss of the plant extracts, especially the essential oils during the incubation period. All the micro-titre plates were then incubated at 37°C for 24 h. After incubation 40 μL (0.04%) of p-iodonitrotetrazolium violet solution (INT) was added to each well of the plate. The plates were allowed to stand for anything between 3 to 6 h depending on the micro-organism to allow for a colour change to occur. The MIC was determined by the lowest dilution which presented with no colour change. The study was done in triplicate on alternate days. Comparative evaluation between the reference and resistance strains were examined and only differences of more than one dilution factor were considered worthy of noting. 3. Results The MIC results for the essential oils as well as the organic and aqueous extracts of the five selected plant species are given in Table 2 (Gram-positive test micro-organisms) and Table 3 (Gram-negative test micro-organisms). Efficacies ranged between noteworthy (where MIC values were below 1.00 mg/mL) to poor activity (N 8.00 mg/mL). The inhibitory concentration of the negative control (acetone solvent) for all strains was N 8.00 mg/mL indicating that the solvent had no influence on the activity. The reference strain (shaded area in Tables 2 and 3) showed in most cases higher susceptibility (0.08–0.63 μg/mL) to the positive control ciprofloxacin than the resistant strains which demonstrated various degrees of resistance (0.08–N2.50 μg/mL). There were some cases (Sa7, Sa8 and Sa9; Ef5 and Ef6; Bc1and Bc2) where the Gram-positive resistant strains demonstrated the same susceptibility pattern to the reference strains. One needs to take into cognisance in

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Table 1 Bacterial strains used in the study.

Micro–organism

Staphylococcus aureus

Resistant strain

Non–resistant control strain

AM, CLO, E, DI, CIP, TE, GN, RIF

Sa1b

MRSA HN00541517 (clinical)

AM, CLOX, E, DI, CIP, TE, GN, RIF

Sa2b

MRSA HN00393616 (clinical)

AM, CLOXE, DI, CIP, TE, GN, RIF

Sa3b

MRSA HN00344510 (clinical)

AM, CLOX, CIP, TE, GN ,RIF

Sa4b

MRSA HN00544873 (clinical)

CLOX, E, DI, CIP, TE, GN

Sa5b

MRSA HN00301932 (clinical)

AM, CLOX, E, CIP, GN, CHL

Sa6b

GMRSA ATCC 33592 (reference)

GN, MET

Sa7c

MRSA ATCC 43300 (reference)

MET

Sa8c

No resistance

Sa9c

ENTFA VRE HN00673964 (clinical)

AM, E, DI, CIP, VA, TP

Ef1b

ENTFE VRE HN00596426 (clinical)

AM, E, DI, VA, TP

Ef2b

ENTFE VRE HN00156099 (clinical)

CIP, TE, VA, TP,

Ef3b

ENTFE VRE HN00109544 (clinical)

CIP, VA, TP

Ef4b

ATCC 51299

VA

Ef5b

No resistance

Ef6c

STREP

Bc1d

No resistance

Bc2c

ESCCO ESBL HN00680299 (clinical)

AM, AMX, FOX, CXM, FEP, CIP, GN

Ec1b

DSM 22314

CAZ

Ec2d

No resistance

Ec3c

AM, AMX, CXM, CTX, CAZ, FEP, CIP, GN AM, AMX, CZ, FOX, CXM, CTX, CAZ, FEP, CIP, GN, MEM, IP, PTZ, TOB, ETP AM, AMX, CXM, CTX ,CAZ, FEP

Kp1b

No resistance

Kp4c

PSEAE HU00738382 (clinical)

AM, CIP, GN, MEM, IP, PTZ, AK

Pa1b

PSEAE HU00743971 (clinical)

Pa2b

ATCC 27853

AM, CAZ, FEP, CIP, GN, MEM, IP, PTZ, AK No resistance

ATCC 13880

AM, AMX, CZ, FOX, CXM, CTX, CAZ, FEP, CIP, GN, MEM, IP, PTZ TOB, ETP No resistance

Sm2c

ACIBA HN00602735 (clinical)

AM, CAZ, FEP, CIP, GN, MEM, IP, PTZ

Ab1b

ACIBA HN00605527 (clinical)

AM, CAZ, FEP, CIP, GN, MEM, IP, PTZ, AK, TOB No resistance

Ab2b

ATCC 29212 Bacillus cereus

DSM 6891 ATCC 11778

Escherichia coli

ATCC 8739 Klebsiella pneumoniae

KLEPN ESL HN00677432 (clinical) KLEPN CRE HN00589933 (clinical)

KLEPN ESBL HN00335335 (clinical) ATCC 13883 Pseudomonas aeruginosa

Serratia marcescens

Acinetobacter baumannii

Code (1–9); Sourceb–d

MRSA HN00562243 (clinical)

ATCC 25923 Enterococcus faecalis

Recorded antibiotic resistance profilesa

SERMA HN00604536 (clinical)

ATCC 19606

Kp2b

Kp3b

Pa3c Sm1b

Ab3c

a

AK = Amikacin; AM = Ampicillin; AMX = Amoxycillin; CAZ = Ceftazidime; CIP = Ciprofloxacin; COL = Colistin; CLOX = Cloxacillin; CHL = Choramphenicol; CTX = Cefotaxime; CAZ = ceftazidime; CXM = Cefuroxime; CZ = Cefazolin; DI = Clindamycin; E = Erythromycin; ETP = Ertapenem; FEP = Cefepime; FD = Fucidin; FOX = Cefoxitin; GN = Gentamycin; IP = Imipenem; MEM= Meropenem; MET = Methicillin; PTZ = Piptaz; RIF = Rifampicin; STREP = Streptomycin; TOB = Tobramycin; TE = Tetracycline; TP = Teicoplanin; VA = Vancomycin. b Clinical strains sourced from NHLS Infection Control and Microbiology Laboratory, University of Witwatersrand. c ATCC strains sourced from Davies Diagnostics. d Deutsche Summlung von Mikrooganismen (DSM) strains; Shaded area = reference strain.

these cases that the resistance patterns (Table 1) may not include ciprofloxacin. 3.1. Resistance patterns towards the Gram-positive test micro-organisms (Table 2) Some variability between the resistant strains of S. aureus (Sa1-Sa8) to that of the reference strain (Sa9) was observed. The organic extracts of the plant species showed the highest variance in response and in all

cases where there was variance, the resistant strains responded better (showed higher efficacy). A similar trend was noted for the essential oils. Only one exception was apparent; the essential oil of O. asteriscoides, with an MIC value of N8 mg/mL against strain Sa8 demonstrated much lower susceptibility than the reference strain (4 mg/mL). Six of the other resistant strains (Sa1-Sa4, Sa6 and Sa7) showed higher susceptibility than the reference strain tested. Four of the resistant strains (Sa1, Sa2, Sa5 and Sa6) responded better to the aqueous extracts of A. afra. The highest variance noted (15 fold) was

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Table 2 The antimicrobial efficacy of a selection of South African medicinal plants against resistant and reference Gram-positive test strains.

Straina

Test sample (MIC mg/ml)

S. aureus

Sa1 Sa2 Sa3 Sa4 Sa5 Sa6 Sa7 Sa8 Sa9 E. faecalis Ef1 Ef2 Ef3 Ef4 Ef5 Ef6 B. cereus Bc1 Bc2

Artemisia afra

Lippia javanica

Control

Croton grattisimus

Osmitopsis asteriscoides

Tetradenia riparia

ORb 0.50 1.00 1.00 1.00 1.00 0.50 0.13 0.13 1.00

AQc 2.00 2.00 >8.00 >8.00 2.00 2.00 >8.00 >8.00 >8.00

EOd 2.00 2.00 2.00 1.00 2.00 1.00 2.00 2.00 4.00

OR 1.00 2.00 1.00 1.00 2.00 0.50 1.00 2.00 2.00

AQ >8.00 >8.00 >8.00 >8.00 >8.00 >8.00 >8.00 >8.00 >8.00

EO 2.00 2.00 2.00 1.00 2.00 1.00 2.00 2.00 4.00

OR 1.00 2.00 1.00 1.00 2.00 0.50 1.00 2.00 2.00

AQ 8.00 >8.00 >8.00 >8.00 >8.00 >8.00 >8.00 >8.00 >8.00

EO 0.50 1.00 1.00 1.00 2.00 1.00 0.50 1.00 2.00

OR 0.25 0.25 1.00 2.00 1.00 0.50 0.50 1.00 2.00

AQ 4.00 4.00 8.00 >8.00 8.00 8.00 8.00 >8.00 4.00

EO 1.00 1.00 1.00 1.00 2.00 1.00 1.00 >8.00 4.00

OR 0.25 0.13 1.00 1.00 2.00 0.50 2.00 2.00 2.00

AQ >8.00 >8.00 >8.00 >8.00 >8.00 >8.00 >8.00 >8.00 >8.00

EO 0.50 1.00 1.00 1.00 2.00 1.00 0.50 1.00 2.00

Positive control (µg/ml) Ciprofloxacin >2.50 >2.50 >2.50 2.50 >2.50 1.25 0.63 0.63 0.63

Negative control

2.00 2.00 4.00 1.00 2.00 1.00

>8.00 >8.00 >8.00 >8.00 >8.00 >8.00

4.00 2.00 2.00 2.00 4.00 4.00

1.00 1.00 2.00 1.00 0.25 2.00

>8.00 >8.00 >8.00 >8.00 >8.00 >8.00

4.00 2.00 2.00 2.00 2.00 4.00

1.00 1.00 2.00 1.00 4.00 2.00

>8.00 >8.00 >8.00 >8.00 >8.00 >8.00

2.00 2.00 2.00 2.00 2.00 4.00

1.00 1.00 1.00 1.00 2.00 2.00

>8.00 >8.00 >8.00 >8.00 >8.00 4.00

4.00 2.00 2.00 2.00 4.00 4.00

1.00 1.00 1.00 2.00 2.00 2.00

>8.00 >8.00 >8.00 >8.00 >8.00 >8.00

4.00 2.00 2.00 2.00 2.00 4.00

2.50 2.50 2.50 0.31 0.63 0.63

>8.00 >8.00 >8.00 >8.00 >8.00 >8.00

0.25 0.50

>8.00 >8.00

1.00 1.00

0.25 0.25

>8.00 >8.00

0.50 0.50

0.25 0.10

>8.00 >8.00

0.13 0.25

0.13 0.50

>8.00 >8.00

0.25 0.50

0.25 0.25

>8.00 >8.00

0.13 0.25

0.08 0.08

>8.00 >8.00

Solvent >8.00 >8.00 >8.00 >8.00 >8.00 >8.00 >8.00 >8.00 >8.00

a

Strain number corresponds to Table 1 Organic extract. c Aqueous extract. d Essential oil; shaded cells = ATCC non-resistant strain; Shaded area = reference strain. b

that where the medicinal plant T. riparia was tested against the resistant strain Sa2 (MIC value of 0.13 mg/mL) compared to the reference strain Sa9 (MIC value of 2.00 mg/mL). Resistance patterns against E. faecalis show little efficacy variation between the resistant strains and the reference strain. One strain (Ef3) showed reduced efficacy (MIC value of 4.00 mg/mL compared to the reference strain Ef6 with an MIC value of 1.00 mg/mL) when tested on the organic extract of A. afra. Another strain (Ef5) showed greater susceptibility (MIC value of 0.25 mg/mL compared to the reference strain Ef6 with an MIC value of 2.00 mg/mL) when tested against the organic extract of L. javanica.

For B. cereus, all plant samples showed similar susceptibility patterns between the resistant strain (Bc1) and the reference ATCC strain (Bc2). One exception was evident where the resistant strain showed higher susceptibility (MIC value of 0.13 mg/mL) when compared to the Bc2 strain (MIC value of 0.50 mg/mL) against the organic extract of O. asteriscoides. 3.2. Resistance patterns towards the Gram-negative test micro-organisms (Table 3) Susceptibility against the E. coli strains showed that the organic extracts of A. afra, O. asteriscoides and T. riparia displayed greater activity

Table 3 The antimicrobial efficacy of a selection of South African medicinal plants against resistant and reference Gram-negative test strains.

E. coli

ORb 0.25 Ec1 2.00 Ec2 2.00 Ec3 K. pneumoniae 0.50 Kp1 1.00 Kp2 1.00 Kp3 4.00 Kp4 P. aeruginosa 1.00 Pa1 1.00 Pa2 S. marcescens 0.25 Sm1 1.00 Sm2 A. baumannii 1.00 Ab1 1.00 Ab2 2.00 Ab3 a

Artemisia afra

Lippia javanica

Croton grattisimus

Osmitopsis asteriscoides

AQc >8.00 >8.00 >8.00

EOd 3.00 4.00 4.00

OR 1.00 2.00 2.00

AQ >8.00 >8.00 >8.00

EO 2.00 4.00 3.00

OR 1.00 2.00 2.00

AQ >8.00 >8.00 >8.00

EO 1.00 2.00 2.00

OR 0.50 2.00 2.00

AQ 8.00 >8.00 >8.00

EO 2.00 2.00 2.00

OR 0.25 2.00 2.00

AQ >8.00 >8.00 >8.00

EO 1.00 2.00 1.00

Positive control (µg/ml) Ciprofloxacin >2.50 1.25 0.32

>8.00 >8.00 >8.00 >8.00

1.00 2.00 1.00 4.00

1.00 1.00 1.00 2.00

>8.00 >8.00 >8.00 >8.00

1.00 2.00 1.00 4.00

1.00 1.00 1.00 1.00

>8.00 >8.00 >8.00 >8.00

1.00 2.00 1.00 2.00

0.25 1.00 0.50 1.00

4.00 >8.00 >8.00 >8.00

1.00 2.00 1.00 2.00

0.25 1.00 0.50 2.00

>8.00 >8.00 >8.00 >8.00

1.00 2.00 1.00 2.00

2.50 2.50 >2.5 0.32

>8.00 >8.00 >8.00 >8.00

>8.00 >8.00

2.00 2.00

0.25 0.50

>8.00 >8.00

2.00 2.00

1.00 1.00

>8.00 >8.00

1.00 2.00

0.25 1.00

8.00 8.00

1.00 2.00

0.25 2.00

>8.00 >8.00

1.00 2.00

>2.5 0.32

>8.00 >8.00

>8.00 >8.00

1.00 1.00

1.00 1.00

>8.00 >8.00

1.00 2.00

1.00 1.00

>8.00 >8.00

1.00 1.00

0.30 1.00

2.00 4.00

1.00 2.00

0.50 2.00

>8.00 >8.00

1.00 2.00

>2.5 0.32

>8.00 >8.00

4.00 >8.00 >8.00

1.00 1.00 2.00

1.00 1.00 1.00

>8.00 >8.00 >8.00

1.00 1.00 2.00

2.00 1.00 1.00

>8.00 >8.00 >8.00

2.00 1.00 2.00

1.00 1.00 1.00

8.00 8.00 >8.00

1.00 1.00 2.00

2.00 1.00 2.00

>8.00 >8.00 >8.00

1.00 1.00 2.00

>2.5 >2.5 0.63

>8.00 >8.00 >8.00

Strain number corresponds to Table 1 Organic extract. c Aqueous extract. d Essential oil; shaded cells = ATCC non-resistant strain; Shaded area = reference strain. b

Tetradenia riparia

Negative control Solvent >8.00 >8.00 >8.00

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against resistant strain Ec1 with an MIC value of 0.25 mg/mL, having higher susceptibility than the reference strain Ec3 (MIC value of 2.00 mg/mL). The resistant strains of K. pneumoniae demonstrated a number of instances where higher susceptibility patterns (lower MIC values) were observed in comparison to the reference strain Kp4. The highest susceptibility difference was noted against the organic extract of T. riparia where the resistant strain Kp1 with an MIC value of 0.25 mg/mL demonstrated an eight fold increased susceptibility compared to the activity of the reference strain Kp3 (MIC value of 2.00 mg/mL). Most of the plant samples showed similar susceptibility patterns between the resistant strain of P. aeruginosa (Pa1) and the reference strain (Pa2). The organic extracts of O. asteriscoides and T. riparia, however, showed higher susceptibilities (MIC values of 0.25 mg/mL) against the resistant strains than the reference strain (MIC values of 1.00 and 200 mg/mL respectively). Similar susceptibility patterns were observed for both the resistant S. marcescens (Sm1) and reference (Sm2) strain. The organic extracts of A. afra, O. asteriscoides and T. riparia responded better (MIC values of 0.25 mg/mL, 0.30 mg/mL and 0.50 mg/mL respectively) to Sm1 than Sm2 (MIC values of 1.00 mg/mL, 1.00 mg/mL and 2.00 mg/mL respectively). All resistant strains of A. baumannii (Ab1 and Ab2) showed similar susceptibility patterns to the reference strain Ab3 with the exception of Ab1 showing increased susceptibility against the aqueous extract of A. afra (MIC 4.00 mg/mL).

4. Discussion All the plant species have been well studied with respect to antimicrobial properties and the results from the reference strains are more or less congruent with literature (Viljoen et al., 2003; Van Vuuren, 2007; Braithwaite et al., 2008; Van Vuuren and Viljoen, 2008; Shikanga et al., 2010; Suliman et al., 2010; York et al., 2012). Results differed where different ATCC strains were used. Also, variations as a result of solvent, geographical location and chemotype were apparent in some cases. Method variation for some plants in the study made comparison of efficacy difficult (Mangena and Muyima, 1999; Shikanga et al., 2010). More importantly it is the analysis of how the resistant clinical pathogens respond to the plant samples compared to the reference strains. In order to analyse this it is important to look at emerging resistance patterns of the pathogens selected in this study and the effects on medicinal plants. Methicillin-resistant S. aureus remains one of the most troublesome resistant pathogens encountered in the healthcare field. With the spread of antibiotic resistance, the treatment of infectious diseases caused by resistant strains of S. aureus is becoming more and more difficult to treat (Lindsay, 2013). In the South African health care setting, the resistance of S. aureus to various antibiotics (cloxacillin 29%, erythromycin 38% and gentamicin 20%) demonstrated the severity of S. aureus resistance (Nyasulu et al., 2012). Although S. aureus is a well-studied micro-organism with respect to South African medicinal plants (Van Vuuren and Holl, submitted for publication), very few studies have investigated activity against resistant strains. A selection of South African medicinal plants against gentamycin and methicillin resistant S. aureus (GMRSA), methicillin resistant S. aureus (MRSA) and a reference strain of S. aureus has been previously investigated (Mabona et al., 2013), however, the study did not include any of the five medicinal plants used in this study. Further comparisons with the antibiotic control were also not made. Also, only reference strains were included in this past study. Clinical strains having been exposed to antibiotics may not be the best strain to use if testing efficacy of plant samples, however, in this study, it is the more appropriate strain as a realistic response to pathogens encountered in a clinical setting is shown.

Enterococci have been shown to have decreased susceptibility to β– lactam antibiotics, particularly cephalosporins and semi-synthetic penicillins. They have also been shown to be tolerant to vancomycin and have been shown to be intrinsically resistant to most common antimicrobial agents, making them the most difficult infectious agents to treat (Miller et al., 2016). Enterococci demonstrating multiple resistance profiles have been isolated from hospital wastewater in Alice (Eastern Cape of South Africa) (Iweriebor et al., 2015). Global studies on natural products tested on resistant E. faecalis strains include the investigation of essential oils in both a planktonic and biofilm state (Benbelaïd et al., 2014) and more recently the synthesis of gold nanoparticles using the aqueous peel extract of Musa paradisiaca (Vijayakumar et al., 2017). Even though some global attention has been given to resistant strains of E. faecalis, no study could be found on South African plants and resistant strains for this pathogen. Most of B. cereus pathogenesis occurs as food poisoning due to undercooked foods. Bacillus cereus confers resistance through formation of biofilms which enables it to produce highly resistant adhesive spores which increases resistance to antimicrobials (Majed et al., 2016). Natural product studies on resistant B. cereus strains both on a global scale and from a South African perspective are lacking and these results comparing the resistant B. cereus strain with the reference strain appears to be novel. Since E. coli colonizes the gastro-intestinal tract of many animals, it may act as a reservoir of resistance genes which may contribute greatly to the spread of resistance. Extended-spectrum beta-lactamases (ESBL) are important as they are easily transferred between different strains by means of plasmids (conjugation) and confer resistance to beta-lactam antibiotics for human and veterinary use (Ben Sallem et al., 2014; Alonso et al., 2016). In a South African study (Abong'o and Momba, 2009), two E. coli isolates were found to be resistant against all eight antibiotics tested. Global natural product studies on resistant E. coli strains include studies on Rhazya stricta, a native herbal shrub of Asia and the investigation on reversal agents from the compound 4-hydroxy-atetralone isolated from Ammannia baccifera (Dwivedi et al., 2014; Khan et al., 2016). This is the first report on E. coli resistant strains tested against South African medicinal plants. Klebsiella pneumoniae is part of the normal flora of the human mouth, gastrointestinal tract and skin (Ryan and Ray, 2004). It is reported to be amongst the most common causes of communityassociated bacterial pneumonia and a number of resistant strains of K. pneumoniae have been identified. Multi-drug resistance has been reported against many antibiotics including trimethoprim/sulfamethoxazole, tetracycline's, aminoglycosides, fluoroquinolones and chloramphenicol (Burgess et al., 2001). There has been an increase in carbapenem-resistant strains of K. pneumoniae which have caused serious challenges to the healthcare sector over the past two decades or so (Anderson et al., 2011). In South Africa, resistance to of K. pneumoniae to ten antibiotics have been noted. These include ciprofloxacillin (35%), cefuroxime (52%), gentamicin (50%) and ampicillin (99%) amongst others (Nyasulu et al., 2012). Even though there has been some natural product studies on resistant strains of K. pneumoniae (Khan et al., 2016; Mishra et al., 2017), no similar studies have previously been undertaken on south African medicinal plants. Pseudomonas aeruginosa is a Gram-negative aerobic bacterium that conveys resistance through efflux pumps and biofilm formation (Poole, 2004). Pseudomonas aeruginosa has been identified as part of the highly resistant micro-organisms commonly termed ESKAPE which includes the following pathogens; Enterococcus faecium, S. aureus, K. pneumoniae, A. baumannii, P. aeruginosa, and Enterobacter species (Rice, 2008). In South Africa the resistance amongst P. aeruginosa isolates has escalated. A recent review described resistance patterns to ciprofloxacillin (43%), gentamicin (50%), amikacin (35%), aztreonam (42%) and even resistance to polymyxin (Nyasulu et al., 2012). Natural products tested against resistant strains of P. aeruginosa include that undertaken on essential oils (Ante et al., 2016) and extracts (Chakraborty

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et al., 2014). No studies other than that reported herein were found on South African medicinal plants tested against resistant strains of P. aeruginosa. Serratia marcescens is a rod-shaped Gram-negative opportunistic pathogen which demonstrates resistance by inclusion of physical or chemical diffusion barriers to antimicrobial agent penetration into the biofilm (Mah and O'Toole, 2001). In South Africa, extensive resistance including resistance to last resort antibiotics such as colistin has been reported for S. marcescens (Sekyere, 2016). Even though S. marcescens resistance is clearly a problem, little, if any attention has been given to natural product studies on these resistant strains. Acinetobacter baumannii is an aerobic Gram-negative bacillus isolated with high frequencies in hospital environments and hospitalized patients (Antunes et al., 2014). Multi-drug resistant A. baumannii has been identified as a rapidly emerging pathogen, causing severe hospital-acquired systemic infections such as pneumonia, bacteraemia, meningitis, wound and urinary tract infections (Maragakis and Perl, 2008). Some strains are resistant to carbapenems or to more than three classes of antimicrobials (Falagas et al., 2006). In South Africa, resistance to A. baumannii has been reported since 2002 (Marais et al., 2004) and is ongoing (Dramowski et al., 2017). Although there is evidence on the antimicrobial properties of natural products when exposed to resistance strains of A. baumannii (Dramowski et al., 2017), no other study except what has been reported herein has been undertaken on South African plants tested against these resistance strains. Looking at the five medicinal plants selected for the study, it is quite clear that in most cases all five plant species respond best to the Grampositive resistant strains of S. aureus than the reference strain. All plants respond differently, but given that the chemistry of the selected plants vary, this is not surprising. Artemisia afra (followed by T. riparia) showed the best susceptibility patterns against the resistant strains of S. aureus and this gives credence to the popularity of A. afra in traditional South African medicine (Liu et al., 2009). For the Gram-negative strains, O. asteriscoides, followed by T. riparia showed the best susceptibility patterns against the resistant strains. 5. Conclusion Most of the clinically resistant strains showed reduced susceptibility to the antibiotic control ciprofloxacin, whereas the reference strains all responded within the expected breakpoint MIC values. In comparison with the plant samples, the reverse was mostly apparent where the resistant strains were more susceptible than the reference strain. The organic extracts followed by the essential oils responded the most favourably to the resistance strains and this is no surprise as both of these samples have multiple compounds which prove to be more difficult for bacteria to convey resistance against. It will be interesting if future studies could be undertaken to determine if plant combinations provide even better response patterns. Each of the five plants responded differently to the resistance strains but overall it is clear that better if not similar susceptibility was observed against the resistant strains in comparison to the corresponding reference strain. The most promising plant species demonstrating better activity against the resistant strains (in order of efficacy) are Artemisia afra followed by T. riparia and O. asteriscoides. Very little progress globally has been made in combatting the onslaught of microbial resistance. This study confirms what has been postulated (but never demonstrated) in many previous studies; medicinal plants respond better to resistant strains that conventional antibiotics. Acknowledgements Mr. Andrew Hankey from the Walter Sisulu botanical gardens is thanked for supplying plant material. Dr. Teena Thomas from NHLS Infection Control and Microbiology Laboratory is thanked for the supply of resistant strains. Mrs. Phumzile Madondo is acknowledged for

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