Hematopoietic Elements in Osteoarthritic Femurs Compared to Normal Bone Marrow as Evaluated by Immunohistochemistry

Introduction

Affecting over 60% of the elderly population, osteoarthritis (OA) has come to be one of the most common joint diseases to affect aging society and leads to the incapacity of joint movement [1]. Caused primarily by mechanical factors including age, obesity, inactive lifestyle, joint injury, and occupation, alterations in gene expression in cartilage and subchondral bone tissue may accelerate the rate of OA disease progression [2]. During alteration of joints in OA, the cartilage covering the ends of the bone become eroded, so the bone surfaces no longer glide across each other, but grind into each other releasing pieces of cartilage and bone into the joint space thereby causing pain and disability. The degenerative changes which include thinning of cartilage layers, increase in subchondral bone content, increased underlying bone remodeling, and fissures which penetrate both cartilage and bone have been well documented [2-9]. Qualitative and quantitative studies have evaluated adipocyte, T-cell, B-cell, macrophage, megakaryocyte, granulocyte, osteoblast, osteoclast, and osteocyte activity that affects the cartilage and subchondral bone in OA [10-18]. The goal of the study was to evaluate the hematopoietic elements in the proximal end of the osteoarthritic femur head as they compare to hematopoietic elements in normal bone marrow aspirates collected from the traditional posterior iliac crest to determine whether the joint pressure and bone degeneration influences the underlying blood cell fractions.

Materials and Methods

This study was approved by the University of Tennessee Health Science Center Internal Review Board IRB number 22-08576-NHSR. Twenty-one formalin-fixed paraffin-embedded osteoarthritic femur head tissues and 13 normal bone marrow biopsy tissues were obtained from the UT shared tissue resource. Only samples evidencing areas of 40% - 60% hematopoietic cells compared with adipocytes were selected for evaluation, and only those selected areas were used for image collection and analysis. Tissue sections were cut at 4 micrometers, were mounted on plus-charged microscope slides, and were placed in a 60°C oven overnight to dry. A hematoxylin and eosin stain was used to visualize the tissue elements and determine appropriate regions of interest.

An iron stain was performed by deparaffinizing tissue sections through several changes of xylene, 1 minute each, and graded alcohols and deionized (DI) water, 10 seconds each. Equal portions of 10% potassium ferrocyanide (424135000, ACROS Organics, NJ, USA) and 20% hydrochloric acid (A144S-212, Fisher Scientific, NJ, USA) were mixed and sections were submersed in this solution for 20 minutes. After a 1-minute tap water rinse, the sections were stained in nuclear fast red solution (STNFRLT, American MasterTech Scientific, Inc., CA, USA) for 5 minutes, rinsed briefly in running tap water, and then were dehydrated through several changes of absolute ethanol and xylene before being cover slipped with a resinous mounting medium.

For immunohistochemistry (IHC) applications, the Bond™ Polymer Refine Detection Kit (DS9800, Leica Biosystems, IL, USA) was used for visualization of the following antibodies: CD3 (103R- 96, Cell Marque/Sigma Millipore, MO, USA), low pH, 1:400, CD20 (ACR3004B, BioCare Medical, CA, USA) high pH, 1:100, CD68 (M0876, Dako/Agilent, CA, USA), high pH, 1:100, Myeloperoxidase light chain (MPO) (sc-365463, Santa Cruz Biotechnology, CA, USA), high pH, 1:500, CD42b (SZ2) (sc-59052, Santa Cruz), high pH, 1:100, and CD71 (NCL-L CD71-309, Leica Biosystems), low pH, 1:80. The tissue sections underwent heat-induced antigen retrieval using a decloaking instrument and the appropriate solution pH as indicated for each antibody above. The IHC steps consisted of the following: A peroxidase block was applied for 5 minutes and then rinsed briefly using tris buffered saline (TBS) 1X Envision™ Flex Wash buffer (DM831, Dako/Agilent) before application of the primary antibody at room temperature for 20 minutes. Thereafter, sections were rinsed with TBS, the kit post primary antibody was applied for 8 minutes, and another TBS rinse followed. The kit polymer reagent was used for 8 minutes followed by TBS and DI water washes before the kit diaminobenzidine reagent was applied for 2 minutes for chromogenic labeling. A kit hematoxylin counterstain was applied for 5 minutes followed by TBS and DI water rinses. The sections were allowed to air dry and were briefly immersed in xylene prior to cover slipping with a resinous mounting medium.

For CD3, T-cell marker, and CD20, B-cell marker, 3 images were obtained using a 40x objective on an Olympus BX45 light microscope (Olympus Corp, Tokyo, Japan) and CellSens software (Olympus). Positively-labeled cells were counted in each field and results reported as an average. For MPO, myeloid cell marker, CD71, red blood cell precursor marker, and CD68, macrophage marker, 3 images were also captured using a 40x objected. However, because positive cell numbers were very large and because the cells adjacent to each other were difficult to reliably count, positivity was evaluated using NIH ImageJ software (https://imagej.nih.gov). The counts represent the amount of brown chromogen present as a percentage of hematopoietic area analyzed. Adipocytes were subtracted from the areas analyzed. Because the kit utilized an amplification step to increase the chromogenic labeling and therefor overall area occupied by each positively-labeled cell, analyzed fraction percentages may exceed 100% when added together. However, with the exception of the myeloid:erythroid, the fraction populations were not compared against each other, but instead were compared only between the OA patient samples and control group samples. Megakaryocyte counts using CD42b were performed on 3 high-powered fields (40x) and iron positivity was graded with a low power (10x) objective using the grading scale 0 - 3 where 0 = no iron present, 1 = minimal iron stores, 2 = moderate iron stores, and 3 = marked iron stores present. For all cell fractions, only areas which were 40% - 60% hematopoietic cellularity were analyzed. Statistical analysis was performed using Microsoft Excel data analysis tool t-Test: Two-Sample Assuming Equal Variances.

Results

B-lymphocytes, T-lymphocytes, and macrophages were found to be slightly increased in OA specimens as compared to control specimens, but the increases were not statistically significant (Table 1, Figures 1 & 2). The myeloid cell component was found to be decreased in OA as compared to that of the control group, but, again, there was no statistical significance to that value. However, the megakaryocyte component was found to be significantly decreased and the CD71+ erythroid fraction was found to be significantly increased in OA cases over the control group (Table 1, Figures 1-3). Because the red blood cell component was increased with no corresponding increase in the myeloid component, the myeloid to erythroid ratio (M:E) in OA cases was decreased (0.98 ± 0.26) compared to the control group (1.66 ± 0.71) and the difference was statistically significant (p-value 0.0005). The presence of iron stores was noted to be distinctly decreased in OA samples (0.16 ± 0.37) as compared to control bone marrow samples (1.08 ± 1.04) with a p-value of 0.001.

Table 1: Hematopoietic cell fractions in osteoarthritis bone marrow compared to normal bone marrow biopsies.

Note: *Measured as percentage of hematopoietic area occupied.

Figure 1: Average number of T-cells, B-cells, and megakaryocytes per high power field (40x) in osteoarthritis bone marrow as compared with control bone marrow. While numbers of T-cells and B-cells present were not significantly different, the number of megakaryocytes in osteoarthritis cases was significantly decreased.

Figure 2: The relative fraction of macrophages, myeloid cells, and erythroid precursor cells given as a percentage of the available hematopoietic cells in osteoarthritis and normal bone marrow. Although the macrophage and myeloid components were not significantly altered, there were more erythroid precursors found in osteoarthritis bone marrow.

Figure 3: Immunohistochemistry assays on bone marrow. Adipocytes (*) occupied 40% - 60% of bone marrow space in the tested samples. CD71 labeling (green arrows) with brown chromogen highlights red blood cell precursors while unlabeled regions (black arrows) identify non-erythroid hematopoietic elements.

a) Control tissue demonstrated approximately 30% positivity while,

b) Osteoarthritis marrow demonstrated approximately 40% positivity. Scale bar = 50 micrometers.

Figure 4: Iron pigment (black arrows) as assessed by Prussian Blue stain for iron, grades 0-3. Scale bar = 50 micrometers.

Discussion

Cartilage growth and maintenance are carefully regulated by appropriate load bearing on the joint. Increase in water content in cartilage softens the matrix allowing for microtears which enlarge with load bearing and eventually lead to flaking of bone and cartilage into the synovial space [1]. Breakdown of cartilage and subchondral bone may allow for exchange of larger-than-normal molecules to influence signaling pathways and increase cross-talk between the normally unconnected compartments of synovium and bone marrow [19]. Studies have shown cellular fractions such as macrophages, osteoblasts, osteocytes, adipocytes, T-cells, B-cells, megakaryocytes, granulocytes, and osteoblasts found in the bone/bone marrow may affect, or be affected by, the cartilage and subchondral bone changes seen in OA joints [10-18].

This study reviewed the quantity of hematopoietic cells found in subchondral bone marrow in OA proximal femurs. Most hematopoietic elements including myeloid cells, macrophages, T-lymphocytes, and B-lymphocytes were similar in quantity to those found in normal bone marrow biopsy specimens taken from the posterior iliac crest. However, the erythroid precursor fraction was increased in OA and there was a corresponding decrease in iron stores. Although excessive iron can give rise to osteoarthritis symptoms [20,21], the relationship between iron deficiency and OA is scarce in the literature. The most common cause of hyperproliferative red blood cell progenitors in bone marrow is increased erythropoietin released in response to hypoxia sensed by the juxtaglomerular apparatus in the kidneys, typically in response to anemia. Hyperproliferation of red blood cells in the bone marrow in conjunction with decreased iron stores is characteristic of iron deficiency anemia. However, lack of access to patient clinical information including hemoglobin levels, serum iron, transferrin, and ferritin precludes a definitive diagnosis. Indeed, a previous studies suggested that OA patients over 70 years are less likely to be anemic than those who have fractured a femur [22]. While bone marrow aspirates collected from the iliac crest are thought to represent the general bone marrow population in the entire organism, it is uncertain whether discrete bone marrow compartments may evidence increases or decreases in hematopoietic lines based on local environment signals. Joints affected by OA are known to produce a myriad of chemokines, proteases, and interleukins among others which could stimulate or suppress one or more hematopoietic lineages [23]. Thus, the increase in erythroid precursors, decrease in megakaryocytes, and decrease in iron stores may reflect the state of the organismal bone marrow. Alternately, the shift in hematopoietic element production may be specifically localized to the femur head due to the mechanical stress related to OA changes. Estimated to make up <1% of the hematopoietic population under normal circumstances, the decrease in megakaryocytes could be directly related to the increased erythroid fraction resulting in reduced space in the bone marrow for megakaryocytes. Additionally, because megakaryocytes and erythroid precursors putatively stem from the same multipotent hematopoietic cell, signals from erythropoietin may drive differentiation from the megakaryocytic pathway and toward the erythrocytic pathway [24].

One limitation of this project pertains to the fact that the methods used in bone decalcification vary and can have adverse effects on subsequent tissue stains [25]. Decalcifying agents containing acid can reduce iron visualization in tissue sections. While both control and OA bone samples underwent acid decalcification with the same reagent, the length of time each was subjected to the acid reagent varied. Larger tissues such as femur heads undergo decalcification for several hours whereas bone marrow biopsies are decalcified for a brief 45 minutes. Because OA bones were subjected to longer decalcification procedure, the possibility that iron may have been leached out of those tissues cannot be excluded. However, the decalcification solution was composed of a weak acid and two of the OA bone samples did evidence clear iron deposits in tissues outside of the hematopoietic compartment suggesting that the decalcification procedures may not have adversely affected the iron stores present.

In conclusion, erythrocyte precursors in the bone marrow were found to be increased in proximal femurs in OA patients as compared to normal bone marrow biopsy specimens taking from the posterior iliac crest of control subjects. The myeloid component was not significantly decreased but nonetheless resulted in a decreased myeloid to erythroid ratio of cell fractions in the bone marrow. The iron stores were lower in OA patients which, when taken with the increase in red blood cell precursors, suggests possible iron deficiency or altered iron metabolism within the proximal femur compartment in persons with OA. Future studies are needed to assess the serum iron, transferrin, ferritin, and complete blood cell count of OA patients to determine anemia status and possible elucidation for the increased red blood cell components

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Antibacterial Potentials of Zapoteca Portoricensis (jacq) H. M. Hernandez on Medically Important Strains of Bacterial

Introduction

Bacterial infections have a large impact on public health Nguyen, et al. [1]. Diseases can range from mild to severe and sometimes can be deadly if left untreated Tamar, et al. [2]. According to report, more than 2.8 million antibiotic resistant bacterial infections occur in the USA each year, and more than 35,000 people die as a result Li [3]. Additionally, according to center for disease control study, the cost to treat multidrugresistant bacterial has been estimated to be $4.6 billion annually Van Duin [4]. Despite the efforts to combat multidrug resistant bacterial with synergistic drugs, the report of multiple drug resistance in medically important strains of bacteria (such as Streptococcus pyrogene, Streptococcus pneumonia, Klebsiella pneumonia, Staphylococcus aureus, Salmonilla typhi and Bacillus subtilis) is still too high Wernli, et al. [5]. The persistent increase in antibiotic resistant strain of bacteria has led to the development of more potent synthetic antibiotics Gajdács [6]. However, these new antibiotics are scarce, costly and so not affordable particularly in developing countries Liu [7]. This therefore, makes compliance to these antibiotics difficult Gajdács [6]. Therefore, there is need for continuous search for new, effective and affordable antimicrobial agents. Local medicinal plants provide a source of new possible antibiotics that may be potent for some of these bacteria Arwa [8]. One of these very important plants is Zapoteca portoricensis Ikeyi [9]. Zapoteca portoricensis is a perennial shrub with a climbing stem which is woody at its base Ikeyi [9]. This plant belongs to the family of Fabeceae and genus Zapoteca, it is commonly called dead awakener (English) Agbafor [10]. However the Yorubas call it Ule, while the Igbos in Nsukka area, where the plant is majorly used by traditional doctors, calls it Azonta Agbo [11]. Zapoteca portoricensis is widely distributed in tropical rain forest, it is found in tropical Africa: Nigeria, Ghana, Togo, and the Caribbean Islands especially British Virgin Island, Dominican Republic, Grenada, Haiti, Jamaica and Puerto Rico Nwodo [12]. The plant has been used traditionally to treat diseases like tonsillitis (sore throat), fever, convulsion, breast engorgement, stomach disorder, purgative, and amenorrhoea (Joshua [13]). Aqueous and alcoholic extracts of the leaf have been reported to be used in the treatment of gastro intestinal disorders, spasmodic and in the treatment of tonsillitis Ukwe [14].

Reported phytochemical analysis of the roots indicated the presence of a number of secondary metabolites such as saponins, resins, glycosides, flavonoids, alkaloids, terpenoids and steroids Ikeyi [9]. Recently, there has been suggestion on the antimicrobial and anti-inflammatory potentials of Zapoteca portoricensis but no scientific investigation has been carried out to verify these claims Esimone [15]. Therefore, this study investigated the antibacterial potentials of Zapoteca portoricensis root extract against some medically important strains of bacteria.

Materials and Methods

Materials

Plant Materials

Field Collection of Plant Material: Plant roots were collected from areas surrounding Nru village, Nsukka. The plant was identified by Mr Alfred Ozioko of the Department of Botany, University of Nigeria, Nsukka. The materials were cleaned of adhering soil and dust in the field by shaking and were dried at room temperature for one week.

Equipment and Instrument: Incubator (Beckman Coulter Co., Indianapolis, Indiana, USA), Oven (WMT-CNC Industrial Co. Ltd., Chizhou, Anhui, China), pH meter (Hanna Instruments, Woonsocket, Rhode Island, USA), Electric balance (Adams Equipment Inc., Oxford, England, UK), Refrigerator, Autoclave, Rotary evaporator (All Thermo-cool Public Ltd Co., Ilupeju, Lagos, Nigeria), Spectrophotometer, (Spectrum Laboratories, Stamford, Connecticut, USA), Infrared spectrometer, Nucear magnetic resonance spectrometer (All Gallenkamp Co., London, England), Whatman No 1 filter paper (Zibo Xinsu Chemical Industry Ltd., Zibo, Shandong, China).

Methods

Extraction Procedure: The roots were washed and air dried at room temperature for two weeks. The dried roots were pulverized using a mechanical grinder and the weight of the root powder was 550g. The powder was macerated in 3.5 litres of absolute ethanol (Figure 1). The mixture was kept for three days under room temperature. Filtration was done using Wattman No. 1 filter paper. The resulting extract was concentrated under fan at room temperature to avoid denaturation of the active ingredients to obtain a semi-solid mass. The extract was stored in a refrigerator at -4°C one week.

Figure 1: Infra-red spectra of ethanol fraction of the plant roots extract.

Fractionation of the Crude Extract: The crude ethanol extract was fractionated using serial solvent fractionation method Jamil, et al. [16]. The dried crude extract weighed 6.5 g. This dried extract was mixed with silica gel (60 GF for column), in the ratio of 1:2 (w:w). The solvents: chloroform, ethyl acetate, acetone and ethanol respectively, were used to wash the mixture. For a particular solvent the washing was done until a colourless filtrate was obtained, then the solvent was changed to another one in the order above, after air drying the mixture. The fractions were concentrated by allowing the solvents to evaporate under room temperature.

Preliminary Phytochemical Analysis: The crude ethanol extract of the roots were subjected to phytochemical analysis according to the method outlined by Harborne [17]. The phytochemical analysis was done to detect the presence of secondary metabolites, such as alkaloids, tannins, saponins, resins, flavonoids, steroids, glycosides and terpenoids.

Test for Alkaloids: A quantity of 0.2 g of the extract was added to 5 ml of 2% hydrochloric acid and heated on a boiling water bath for 3 min, it was allowed to cool and then filtered. A portion of the filtrate (1ml) was treated with 2 drops of Dragendorff’s reagent, the appearance of a red precipitate indicated the presence of alkaloids. Test for Glycosides: Two gram of the extract was mixed with 30 ml of water and heated on a water bath for 5 min and filtered. About 5 ml of a mixture of equal parts of Fehling’s solutions A and B was added to 5 ml of the filtrate until it turned alkaline (tested with litmus paper) and then boiled on a water bath for 5 minutes. A brick-red precipitate indicated the presence of glycosides.

Test for Flavonoids: Ethyl acetate, 10 ml, was added to 0.2 g of the sample and heated on a water bath for 3 minutes. The mixture was allowed to cool and filtered. A volume of 4 ml of the filtrate was shaken with 1 ml of dilute ammonia solution. The layers were allowed to separate. Absence of a yellow colour in the ammoniacal layer indicated the absence of flavonoids.

Test for Resins: A quantity of 0.2 g of the sample was extracted with 15 ml of 96% ethanol. The alcoholic extract was then poured into 20 ml of distilled water in a beaker. A precipitate occurring indicated the presence of resins.

Test for Tannins: A quantity, 2 g of the sample was boiled with 5 ml of 45% ethanol for 5 minutes. The mixture was cooled and then filtered; the filtrate was then treated with lead sub-acetate solution. To 1 ml of the filtrate 3 drops of lead sub acetate solution was added. A none-red gelatinous precipitate indicated the absence of tannins.

Test for Saponins: A quantity of 0.1 g of the extract was boiled with 5 ml of distilled water on a water bath for 5 min. The mixture was filtered hot and allowed to cool. To 1 ml of the filtrate, 2 drops of olive oil was added and the mixture shaken vigorously. The formation of emulsion indicated the presence of saponins.

Figure 2: 13C NMR spectra of ethanol fraction of the plant roots extract.

Test for Terpenoids and Steroids: A volume (9 ml) of absolute ethanol was added to l g of sample and refluxed for 5 min and filtered. The filtrate was concentrated to 2.5 ml on a boiling water bath and then 5 ml of hot water was added. The mixture was allowed to stand for 1 hr and the waxy matter was filtered off. The filtrate was extracted with 2.5 ml of chloroform using a separating funnel (Figure 2). To 0.5 ml of the chloroform extract in a test tube, 1 ml of concentrated sulphuric acid was carefully added to form a lower layer. A reddish brown interface showed the presence of steroids. Another 0.5 ml of the chloroform extract was evaporated to dryness on a water bath and heated with 3 ml of concentrated sulphuric acid for 10 min on a water bath. A grey colour indicated the presence terpenoids.

Test for Acidic Compounds: A quantity, 0.l g was placed in a clean dry test tube and sufficient water added. This was warmed in a hot water bath and then cooled. A piece of moist litmus paper was dipped into the filtrate and the colour change on the litmus paper was observed. A red coloration of the litmus paper indicated the presence of acidic compounds.

Test for Proteins: A volume, 5 ml of distilled water was added to 0.1 g of the extract. This was left to stand for 3 hrs and then filtered. To 2 ml portion of the filtrate was added 0.1 ml Million’s reagent. It was shaken and kept for observation. A yellow precipitate indicated the presence of proteins.

Test for Carbohydrate: A quantity, 0.1 g of the extract was shaken vigorously with water and then filtered. To the aqueous filtrate was added few drops of Molisch reagent, followed by vigorous shaking again. Then, 1 ml of concentrated sulphuric acid was carefully added through the side of the test tube to form a layer below the aqueous solution. A brown ring at the interface indicates the presence of carbohydrate.

Test for Reducing Sugar: A quantity, 0.1 g of extract was shaken vigorously with 5ml of distilled water and filtered. Equal volumes of Fehling’s solutions A and B were added to 1 ml portion of the filtrate. The mixture was shaken vigorously. A brick red precipitate indicated the presence of reducing sugars.

Test for Fats and Oil: A quantity of 0.1 g of sample was pressed between filter paper and the paper observed. A control was also prepared by placing 2 drops of olive oil on filter paper. Translucency of the filter paper indicates the presence of fats and oil.

Test microorganisms: The clinical isolates were obtained from the Faculty of Veterinary Medicine, University of Nigeria, Nsukka, Enugu State, Nigeria. Test isolates were Streptococcus pyogenes, Streptococcus pneumonia, Bacillus subtilis, Salmonella typhi, Staphylococcus aureus, E. coli, Klebsiella pneumonia.

Preparation of culture media: The materials for the experiment were sterilized in an autoclave. They were allowed to cool before being used. Nutrient agar, pH 7.4 (Oxoid laboratories, England, United Kingdom) was used in the study. A quantity, 5.6 g of dried nutrient agar was weighed. This was dissolved in 200 ml of water. The suspension was well mixed by stirring in the water. The solution was heated to dissolve and to purify media. The media were sterilized by autoclaving at 50°C. The plates were thereafter inoculated with test microorganisms and incubated at 37°C for 24 hours.

Determination of antimicrobial activity: The antimicrobial activities of the crude extract were determined using NCCLS method (WHO [18]). For determination of antibacterial activity, bacterial cultures were adjusted to 0.5 McFarland turbidity standards and inoculated to 15 cm diameter nutrient agar plates. The crude ethanol extract of the plant’s roots was dissolved in dimethyl sulfoxide (DMSO); (500 mg of the extract was constituted with 5 ml of 100% DMSO to prepare stock solution). The concentration of stock was 100 mg/ml. Different concentrations (50 mg/ml, 25 mg/ml, 12.5 mg/ml, 6.25 mg/ml), of the plant root extract were prepared using serial dilution in DMSO. The control had DMSO alone without any extract. The method used was agar well diffusion method (Arwa, et al. [8]). Briefly, microorganisms from growth on nutrient agar incubated at 37°C for 18 h were suspended in saline solution, 0.85% NaCl and adjusted to a turbidity of 0.5 Mac Farland standards (108 cfu/ml). The suspension was used to inoculate 15cm diameter Petri plates with a sterile non-toxic cotton swab on a wooden applicator. Six millimeter diameter wells were punched in the agar and filled with fractions. The dissolution of the extract was aided by 1% (v/v) DMSO which did not affect microorganisms’ growth, according to our control experiments. Commercial antibiotic, streptomycin was used as positive reference standard to determine the sensitivity of the strains. The extract was introduced into the wells. Plates were incubated in at 37°C for 24 hr. Antibacterial activities were evaluated by measuring inhibition zone diameters. The experiments were conducted twice. All tests were performed in duplicate and the antibacterial activity was expressed as the mean diameter of inhibition zones (mm) produced by the plant root extracts.

Use of Infra-Red and NMR Spectroscopy to Determine the Functional Groups in the Ethanol Fraction

Infrared spectroscopy was carried out on the ethanol fraction to determine the functional group present in the fraction. The extract was dissolved in ethanol and ethanol was used as a control. Infrared spectra were obtained between the frequencies: 399.19 to 3999.64, characteristic peak of the functional group were detected and shown in (Table 5). The NMR spectroscopy was carried out on the samples, using DMSO as the solvent and teramethyl silane as an internal standard, at frequency of 199.9671071MHz at ambient temperature. The characteristic peaks of the functional groups were detected and shown in (Table 6).

Result

Extraction and Fractionation

The percent extract yield of the plant roots was 1.56%. After fractionation the following fractions were obtained: chloroform fraction, acetone fraction, ethyl acetate fraction and ethanol fraction.

Result of Phytochemical Analysis and Macronutrients in the Crude Extract of the Plant Roots

Phytochemical screening of the plant roots showed that the plant contains alkaloids, glycosides, reducing sugar, carbohydrates, steroids, terpenoids, saponins, proteins, and fats and oil. The roots lacked flavonoids, tannins and acidic compounds. The chemical classes of compounds in the plant roots as indicated by the phytochemical analysis are listed in (Table 1).

Table 1: Phytochemical constituents of the plant.

Note: Absent -, Present, Low concentration ++ Moderate concentration +++ High concentration ++++.

The Antimicrobial Activities of the Crude Ethanol Extract

Table 2: Inhibitory zone diameter of different concentration of the extract.

(Table 2) shows the inhibition zone diameter (IZD) of the crude ethanol extract on the microorganisms used. The crude ethanol extract of the plant roots was found to possess antimicrobial activities on both Gram-positive and Gram-negative bacteria. The crude extract was active on S. Pyogenes, Streptococcus pneumonia, Klebsiella pneumonia, Staphylococcus aureus, Salmonella typhi and E. coli. The crude extract was not active on Bacillus subtilis. There were significant differences (p<0.05) between the standard antibiotics and crude extract.

The Minimum Inhibitory Concentrations (Mic) of the Crude Ethanol

MICs of the crude extract are 25 mg/ml, 100 mg/ml, 6.25 mg/ml, 50 mg/ml, 25 mg/ml 6.25 mg/ml for Staphylococcus aureus, Streptococcus pneumonia, Streptococcus pyogenes, E. coli, Klebsiella pneumonia and Salmonella typhi respectively (Table 3).

Table 3: Minimum inhibitory concentration (MIC) of the crude ethanol extract.

Antimicrobial Activities of the Fractions

The chloroform fraction was active on Staphylococcus aureus and on Streptococcus pneumonia, but it was not active on other microorganisms. The ethyl acetate fraction was active only on Streptococcus pneumonia while acetone fraction did not show any significant activity on the microorganisms used. The ethanol fraction showed significant activity on all the bacteria except Bacillus subtilis. Only ethanol fraction of the crude extract was active on both Gram-negative and Gram-positive bacteria (Table 4).

Table 4: Inhibitory zone diameter of the fractions at 100mg/ml.

Infra-Red and Nuclear Magnetic Resonance Spectra

When scanned in infra-red, the H1 at 199.9671071 MHz was observed and the highest peaks were between 2-6 ppm, the following groups present are shown in (Table 5). The 13CNMR showed peaks between 180-56 ppm. It was noticed that the following groups are present in the fraction; aromatic group, sulfuryl group, halogen group, carboxyl group and some double and triple bonded carbons.

Table 5: The suspected functional groups from infrared spectra.

Table 6: The functional group from NMR spectral.

Discussion

This research was carried out to determine the antimicrobial potentials of Zapoteca portoricensis root extract on streptococcus pyrogen and other bacteria. The result show that the root extracts of Zapoteca portoricensis had antimicrobial effect on streptococcus pyrogen, E. coli, Staphylococcus aereus, Klebsiella pneumonia and Salmonilla typhi except Bacillus subtilis. The reason for the antimicrobial effect of the root extract of Zapoteca portoricensis might be because it possesses phytochemical compounds that might have antimicrobial properties (Ikeyi [19]). This is quite true because the phytochemical of the root extract of the plant showed the presence of phytochemicals such as alkaloids, glycosides, reducing sugar, terpenoids, saponin and resins which have been reported to have antimicrobial activities and could serve as scaffolds for important antibacterial drugs (Ikeyi [19]). Specifically, among the phytochemical compound found in the root extract of Zapoteca portoricensis is alkaloids (Agbafor [10]). Alkaloids have been suggested to have good antimicrobial activity against both gram-negative and gram-positive bacterial (Cushnie, et al. [20]). The presence of alkaloids in high concentration could have likely contributed to the antimicrobial activity of the ethanolic root extract of Zapoteca portoricensis. The phytochemical components seen in these study corroborates with the results of (Agbafor [10]) who also confirmed the presence of alkaloids, glycosides, terpenoids and saponin in the root extract of Zapoteca portoricensis. In order to determine the level of antimicrobial activity of the root extract of Zapoteca portoricensis on the bacterial tested, the crude extract of the plant was used to determine the inhibitory zone diameter of the plant at different concentrations. The result showed that the zone of inhibition of the different bacterial by the plant extract were concentration dependent and was highest at the concentration of 100 mg/ml. Among all the bacterial tested, the inhibitory zone diameter of the plant extract on streptococcus pyrogen at the concentration of 100 mg/ml was found to be the highest. This suggests that streptococcus pyrogen has higher sensitivity to the root extract of Zapoteca portoricensis than other bacterial tested. The variations in the sensitivity of the bacterial to the root extract of Zapoteca portoricensis could be due to differences in the cell wall composition of the bacterial (Nguyen, [1]).

Streptococcus pyrogen might possess a cell wall that allows for easy permeability of the chemical constituents of Zapoteca portoricensis root extract and thus allowing for the inhibition of the cell wall synthesis of the bacterial (Gajdács [6]). It has also been suggested that the cell wall of Streptococcus pyrogen lack cell membrane, an attribute that could aid in easy uptake of antimicrobial phytochemicals in Zapoteca portoricensis (since there will be no selective permeability) (Tamar, et al. [2]). This result corroborates with (Agbafor [21]) who reported high inhibitory zone diameter of Streptococcus pyrogenes, Staphylococcus aureus, E. coli and Klebsiella pnuemonia by root extract of Zapoteca portoricensis. Generally, the leaf extract of Zapoteca portoricensis had inhibitory effect on all the bacterial tested except Bacillus subtilis. The mechanism of inhibition of these bacterial could vary, such as inhibition of the protein synthesis, disruption of membrane structure, inhibition of folate coenzymes, inhibition of nucleic acids, inhibition of peptidoglycans, deterioration of cytoplasmic pH, increase permeability of plasma membrane, prevention of extracellular and intracellular microbial enzyme production, interruption of bacterial metabolic pathways and disruption of plaque and biofilm formation (Mahon, et al. [22]) (Lin, et al. [23]). Depending on the morphological and physiological structure of the bacterial, the demonstration of antimicrobial activity against both gram-positive and gram negative bacteria by Zapoteca portoricensis may be indicative of the presence of broad spectrum antibiotic compounds in the extract (Reygaert [24]). Different fractions of the leaf extract of Zapoteca portoricensis were assayed in order to determine which fraction has higher antimicrobial activity. The result showed that only the ethanol fraction had significant activity on both the gram-negative and gram-positive bacteria. This might be due to better solubility of the active component of Zapoteca portoricensis in ethanol and suggests that ethanol might be the best solvent of extraction of antimicrobial compounds in Zapoteca portoricensis (Nwodo, et al. [25]). This corroborates with the result of (Nwodo, et al. [25]) who reported better solubility of Zapoteca portoricensis in ethanol than other solvents. Next, investigation was carried out to determine the minimum inhibitory concentration (MIC) of the root extract of Zapoteca portoricensis on the bacterial.

The result showed that Streptococcus pyrogenes and Salmonella typhi had the lowest MIC scores indicating that less concentration of ethanolic root extract of Zapoteca portoricensis is required for inhibition of these bacterial (Nkechukwu, et al. [26]). This suggests that, ethanolic root extract of Zapoteca portoricensis might be a more effective antimicrobial agent on Streptococcus pyrogenes and Salmonella typhi than other bacterial tested. The efficacy of phytochemicals in the plant extracts as antimicrobial agents with little or no side effect depends on the structure of the compounds interacting with the pathogen. The infrared spectrum of the ethanolic root fraction of Zapoteca portoricensis showed a broad band at 3400cm-1, 2900cm-1, 2100cm-1, 1600cm-1, 1000cm- 1, 1400cm-1 and 900cm-1 indicating the presence of hydroxyl, aldehyde, cyanide, carbonyl, primary amine, alkene and aromatic group respectively. In addition, the 13CNMR spectra showed peaks between 60-80 ppm and 100-110 ppm indicating the presence of sulfulryl, carboxyl, halide, amide, alkene and aromatic groups. The NMR spectra corroborates with the infrared spectrum which also indicated the presence of amine, alkene and aromatic groups. Earlier studies have shown that functional groups such as alkene, aldehylde, carbonyl, carboxyl and aromatic groups exerts good antibacterial activity and play an important role in antibiotic action (Siyanbola, et al. [27]) (Jabamalairaj, et al. [28]). Since the infrared spectra and NMR showed the presence of these functional groups in ethanolic root fraction of Zapoteca portoricensis [28]. It suggests that these functional groups possess great inhibitory effect on the bacteria tested and could be responsible for the inhibition shown by Zapoteca portoricensis root extract.

Conclusion

In this work, the antimicrobial potentials of Zapoteca portoricensis root extract was evaluated, it was found that ethanolic root extract of Zapoteca portoricensis had inhibitory effect on Streptococcus pyrogene, Staphylococcus aereus, E. coli, Klebsiella pneumonia and Salmonella typhi except Bacillus subtilis. Infrared and NMR spectra of the root extract also showed the presence of functional groups such as alkene, aldehylde, carbonyl, carboxyl and aromatic groups which might have played a significant role in the inhibitory effects of Zapoteca portoricensis on streptococcus pyrogene and other bacterial tested.

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