Quinoline alkaloids from Lunasia amara inhibit Mycobacterium tuberculosis H37Rv in vitro

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References [1] Scholl L, Chang E, Reitz B, Chang J. Sternal osteomyelitis: use of vacuum-assisted closure device as an adjunct to definitive closure with sternectomy and muscle flap reconstruction. J Card Surg 2004;19:453–61. [2] Bouza E, Munoz P. Micro-organisms responsible for osteo-articular infections. Baillieres Best Pract Res Clin Rheumatol 1999;13:21–35. [3] Bain KT, Wittbrodt ET. Linezolid for the treatment of resistant Grampositive cocci. Ann Pharmacother 2001;35:566–75. [4] Lovering AM, Zhang J, Bannister GC, et al. Penetration of linezolid into bone, fat, muscle and haematoma of patients undergoing routine hip replacement. J Antimicrob Chemother 2002;50:73–7. [5] Rana B, Butcher I, Grigoris P, Murnaghan C, Seaton RA, Tobin CM. Linezolid penetration into osteo-articular tissues. J Antimicrob Chemother 2002;50:747–50. [6] Kutscha-Lissberg F, Hebler U, Muhr G, Koller M. Linezolid penetration into bone and joint tissues infected with methicillin-resistant staphylococci. Antimicrob Agents Chemother 2003;47:3964–6. [7] MacGowan AP. Pharmacokinetic and pharmacodynamic profile of linezolid in healthy volunteers and patients with Gram-positive infections. J Antimicrob Chemother 2003;51(Suppl. 2):ii17–25.

Simeon Metallidis ∗ John Nikolaidis Georgia Lazaraki Eleni Koumentaki Vasiliki Gogou Dimitrios Topsis Pavlos Nikolaidis 1st Internal Medicine Department, Infectious Diseases Division, AHEPA University Hospital, Thessaloniki, Greece Nikolaos Charokopos Cardiosurgical Department, AHEPA University Hospital, Thessaloniki, Greece Georgios Theodoridis Chemistry Department, Aristotle University of Thessaloniki, Greece ∗ Corresponding

author. Tel.: +30 6944 361 931; fax: +30 231 099 4615. E-mail address: [email protected] (S. Metallidis) 10 January 2007

doi: 10.1016/j.ijantimicag.2007.01.012

Quinoline alkaloids from Lunasia amara inhibit Mycobacterium tuberculosis H37 Rv in vitro Sir, Tuberculosis (TB) ranks second among the leading infectious diseases in the world, with ca. 2–3 million deaths in 7–8 million new cases of active TB each year. This situation has been aggravated by the emergence of multidrugresistant strains of the TB organism coupled with human immunodeficiency virus (HIV)-induced immunodepression

[1]. Therefore, new classes of potent antimycobacterial drugs, including natural products, are needed to address this problem. It has been reported that several secondary metabolites of plant origin exhibit interesting inhibitory activity against several species of mycobacteria [2,3]. A previous microbiological study on the alcoholic extract of Lunasia amara Blanco (Rutaceae) revealed interesting activity against Mycobacterium smegmatis ATCC 607, Bacillus subtilis ATCC 6633, Staphylococcus aureus ATCC 25923, Mycobacterium tuberculosis H37 Rv and Mycobacterium avium [4,5]. In this communication, we report the inhibitory activity against M. tuberculosis H37 Rv of the quinoline alkaloids obtained from the leaves of L. amara through bioassayguided isolation. We also report the activity of the crude extract, semicrude extracts and fractions from which the alkaloids were isolated. Air-dried leaf samples of L. amara (1 kg) were extracted with ethanol to give a green syrup (275 g). A portion of the crude ethanolic extract (225 g) was subjected to group separation by solvent partitioning using n-hexane (10 g), dichloromethane (17 g), ethyl acetate (5 g) and nbutanol (17 g). The most active hexane and dichloromethane extracts (Table 1) were subjected to further purification. Ten grams of hexane extract was purified by silica gel vacuum liquid chromatography using a step gradient addition of dichloromethane in n-hexane to give four fractions. The most bioactive subfraction (H4) was column chromatographed using silica gel (hexane–ethyl acetate/ethyl acetate–methanol, 10% increments) followed by SephadexTM LH-20 (chloroform) to give 4-methoxy-2phenylquinoline (1) (25.8 mg) (Fig. 1). The dichloromethane extract (17 g) was subjected to silica gel vacuum liquid chromatography (VLC) using dichloromethane in hexane (2:3, 4:1, 1:0, v/v) and methanol in dichloromethane (1:9, 2:8, 4:6, 6:4, 8:2, v/v) to afford six fractions. The most active fraction (D4) was subjected to silica gel column chromatography using hexane–dichloromethane (1:1, 3:2, 4:1, 1:0, v/v) and dichloromethane–methanol (1:9, 7:3, v/v) to give kokusagine or 7,8-methylenedioxydictamine (2) (568.3 mg) (Fig. 1). Another aliquot of the ethanolic extract (50 g) was subjected to acid–base extraction using acetic acid–ammonium hydroxide. The crude alkaloid extract (2.9 g) obtained at pH 4 was purified by VLC in increasing amounts of dichloromethane in hexane (1:1, 3:1, 1:0, v/v) and methanol in dichloromethane (1:3, 1:1, 1:0, v/v) to give four fractions. Fraction 3 (1.4 g) was further purified by VLC and silica gel column using the same solvent to give graveolinine or 4-methoxy-2-(3 ,4 -methylenedioxy)phenyl quinoline (3) (11.7 mg) (Fig. 1). The quinoline alkaloids 1–3 were identified by comparison of their physicochemical constants and spectral data (ultraviolet, infrared, mass, and 1 H and 13 C nuclear magnetic resonance spectra) with the literature [6–8]. The extracts, fractions and isolates were screened for inhibitory activity against M. tuberculosis H37 Rv using a radiorespirometric assay, as described previously [9].

Letters to the Editor / International Journal of Antimicrobial Agents 29 (2007) 738–748 Table 1 Percent inhibitory activities of the various extracts and fractions against Mycobacterium tuberculosis H37 Rv Test concentration

Crude extract Ethanol

1000 ␮g/mL

100 ␮g/mL

99

40 Test concentration

Semicrude extracts n-Hexane DCM Ethyl acetate n-Butanol

100 ␮g/mL

50 ␮g/mL

100 93 −2 8

78 33 0 7

Test concentration 100 ␮g/mL

33 ␮g/mL

Fractions Hexane fractions H1 H2 H3 H4

72 24 97 99

22 16 60 98

DCM fractions D1 D2 D3 D4 D5 D6

18 99 99 99 99 5

17 96 71 93 64 2

DCM, dichloromethane.

Extracts, fractions and isolates (1–3) were solubilised at 10.24 mg/mL in dimethyl sulphoxide (DMSO), filter sterilised and stored at −80 ◦ C until use. Subsequent dilutions were performed in DMSO. Fifty microlitres of solution were added to 4 mL BACTEC 12B broth (Becton Dickinson, Towson, MD) to achieve the desired final concentrations.

Fig. 1. Structures of the isolated quinoline alkaloids from the leaves of Lunasia amara.

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Drug susceptibility testing was performed in BACTEC 460. The minimum inhibitory concentration (MIC) of the isolate was defined as the lowest concentration of test sample that effected a daily growth index (GI) increase and final GI lower than the 1:100-diluted control vial readings when 1:100 GI was >30. For comparison purposes, rifampicin was used as a positive standard (98% inhibition at 0.125 ␮g/mL). Alkaloids 1 and 3 showed more potent inhibitory activity (MIC = 16 ␮g/mL against the test organism) compared with alkaloid 2 (MIC = 33 ␮g/mL). For comparison purposes, rifampicin was used as the positive drug standard (MIC = 0.125 ␮g/mL). Seemingly, it was deduced that the presence of an aryl group, which in this case is a phenyl or a methylenedioxyphenyl ring at the C2 position of the quinoline nucleus, enhances inhibitory activity. Another common structural feature among the three alkaloids that also causes increased bioactivity is the presence of a 4-methoxy group and a fully aromatised quinoline nucleus [3,8]. This study reports for the first time the potential use of alkaloids 1–3 as agents against M. tuberculosis. Moreover, it also extends the structure–activity relationship analysis of antimycobacterial quinoline alkaloids isolated and purified from Rutaceae plants [8,10].

Acknowledgments The authors thank the National Research Council of the Philippines and the International Foundation for Science (IFS) for the financial support, as well as the DOST-ESEP for the scholarship grant to V. Dalangin. References [1] World Health Organization. Tuberculosis. Fact Sheet No. 104. Geneva, Switzerland: WHO; 2004. [2] Okunade AL, Elvin-Lewis MPF, Lewis WH. Natural antimycobacterial metabolites: current status. Phytochemistry 2004;65:1017–32. [3] Copp BR. Antimycobacterial natural products. Nat Prod Rep 2003;20:535–57. [4] Aguinaldo AM, Chua NM. Philippine medicinal plants active against Mycobacterium 607. Acta Manila Ser A 1988;37:81–4. [5] Dalangin VM, Abe F, Yamauchi T, Byrne LT, Franzblau SG, Aguinaldo AM. Transactions of the National Academy of Science and Technology, Philippines. Bicutan, Taguig, Metro Manila: NAST, DOST; 1997. [6] Goodwin S, Smith AF, Velasquez AA, Horning EC. Alkaloids of Lunasia amara Blanco, Isolation studies. J Am Chem Soc 1959;81:6209–13. [7] Goodwin S, Smith AF, Horning EC. Alkaloids of Lunasia amara, 4methoxy-2-phenylquinoline. J Am Chem Soc 1957;79:2239–41. [8] Houghton PJ, Woldemariam TZ, Watanabe Y, Yates M. Activity against Mycobacterium tuberculosis of the alkaloid constituents of Angostura bark, Galipea officinalis. Planta Med 1999;65:250–4. [9] Fischer NH, Lu T, Cantrell CL, Casta˜neda-Acosta J, Quijano L, Franzblau SG. Antimycobacterial evaluation of germacranolides. Phytochemistry 1998;49:559–64. [10] Adams M, Wube AA, Bucar F, Bauer R, Kunert O, Haslinger E. Quinolone alkaloids from Evodia rutaecarpa: a potent new group of antimycobacterial compounds. Int J Antimicrob Agents 2005;26: 262–4.

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Alicia M. Aguinaldo ∗ Victoria M. Dalangin-Mallari Allan Patrick G. Macabeo Phytochemistry Laboratory, Research Center for the Natural Sciences, Thomas Aquinas Research Complex, University of Santo Tomas, Espana, Manila 1008, The Philippines Lindsay T. Byrne School of Biomedical and Chemical Sciences, University of Western Australia, 35 Stirling Highway, Crawley WA 6009, Australia Fumiko Abe Tatsuo Yamauchi Faculty of Pharmaceutical Sciences, Fukuoka University, Nanakuma 8-19-1, Jonan-ku Fukuoka 814-80, Japan Scott G. Franzblau Institute for Tuberculosis Research, College of Pharmacy, University of Illinois at Chicago, 833 S. Wood St., Chicago, IL 60612-7231, USA ∗ Corresponding

author. Tel.: +63 2 406 1611x8224.

E-mail addresses: [email protected], alicia [email protected] (A.M. Aguinaldo) 19 January 2007

doi: 10.1016/j.ijantimicag.2007.02.004

Bactericidal properties of silica particles with silver islands located on the surface Sir, The bactericidal properties of silver have been known for centuries. It was employed in the colloidal form (on the basis of, e.g., silver nitrate) owing to its good antibacterial activity. Modification of various inorganic compounds by addition of silver is an attractive method to protect humans from diseases caused by microorganisms. Nanosized silver particles have a high surface area but have a tendency to form aggregates. Antibacterial activity increases with an increase in silver concentration and decreases with an increase in particle size of silver nanoparticles [1]. Silver nanoparticles with larger diameters have correspondingly lower surface areas. One of the ways of obtaining well-dispersed silver powders is to use them as dopants in silica materials [2–4]. The interaction of silver nanoparticles with organic functional groups of silane compounds prevents the nanoparticles from aggregating [2]. Silica displays good bioactivity and biocompatibility [5]. Silver-doped silica materials are difficult to prepare by the conventional melt–quenching methods but are easy to obtain by the sol–gel method. Silica materials containing silver show high chemical stability and durability.

In this paper, we discuss the bacteriostatic activity of silica particles with nanosilver on the surface. Silver particles have a size of ca. 30 nm and are deposited on silica spheres in the form of islands. The antibacterial effect of these particles was tested on Gram-negative Escherichia coli ATCC 11229 and Gram-positive Staphylococcus aureus ATCC 6538 strains. The uniform, submicron-sized silica particles were synthesised following the base-catalysed polycondensation of tetraethoxysilane in alcoholic medium [6,7]. The prepared particles were characterised by scanning electron microscopy on a Joel JSM 5800LV microscope. To examine the bactericidal effect of the SiO2 –Ag powders on the growth of bacteria, ca. 105 colony-forming units (CFU) of E. coli and S. aureus were cultured overnight in 10 mL of nutrient broth for 18 h at 37 ◦ C. The overnight bacterial cultures were diluted 1000 times, transferred into fresh nutrient broth and were used in further bacterial testing. The samples were incubated for 6 h at 37 ◦ C in a water-bath and 100 ␮L aliquots were collected every hour and plated on Petri dishes with sterilised nutrient agar. The Petri dishes were incubated for 24 h at 37 ◦ C and colonies were counted after 24 h. The counts from three plates corresponding to each sample were averaged. The number of CFU at zero time was taken as 100%. The obtained silica particles have a spherical shape with a diameter of ca. 500 nm. Silver islands have been deposited on the surface of the silica particles. The silver islands are ca. 30 nm in size. Fig. 1 shows the silica particles with the surface silver islands obtained by the sol–gel method. The results of bacterial growth kinetics in the presence of immobile silver on silica spheres are given in Table 1. Colonies (CFU/mL) of S. aureus and E. coli grew ca. 40% over 6 h incubation for the control specimens K1 and K2. Neither S. aureus nor E. coli demonstrated sensitivity to SiO2 . In the presence of 0.5 mg/mL SiO2 –Ag in nutrient broth, the number of S. aureus bacterial cells decreased slowly during the first 3 h of incubation. After 4.5 h of incubation, a decrease of S. aureus from 51 × 105 CFU/mL to 20 × 104 CFU/mL was observed. In the case of E. coli, it appears that the strain was more sensitive to the silica spheres with the silver islands on the surface (0.5 mg/mL; P1 sample) than S. aureus strains. At 6 h, the number of E. coli decreased by 57% with regard to the initial value for the sample P1. SiO2 –Ag inhibited the growth of both bacterial strains. We demonstrated that S. aureus ATCC 6538 and E. coli ATCC 11229 showed sensitivity to the silver on silica spheres (Ag–SiO2 ), irrespective of the structure of their cell walls. In general, Gram-negative bacteria are more susceptible to silver ions than Gram-positive bacteria, owing to the thin peptidoglycan (murein) walls of Gram-negative bacteria (2–3 nm) compared with the thickness of peptidoglycan in Gram-positive bacteria (10 nm). The bacteriostatic effect of the silver ion is induced by the Ag+ adsorption processes to murein. In our experiments, the silica spheres with silver nanoislands inhibited