Materials and Methods3.1. Materials3.
1.1 Chemicals and DrugsChemicals and solvents of laboratory and analytical grade were used throughout this work: Carrageenan (Tokyo chemical industries, Japan), methanol, n-butanol, chloroform, acetylsalicylic acid (Addis Pharmaceutical Factory/APF), acetic acid, morphine (AMINO Ltd. Neuenhof, Switzerland), formalin, distilled water and normal saline were used. 3.
- Thesis Statement
- Structure and Outline
- Voice and Grammar
1.2. Collection, Identification and Preparation of Plant Materials The roots of C. ficifolius were collected from Kilte Awlaelo district of Tigray Regional State, 47.7 kilometers from Mekelle and 829 kilometers from Addis Ababa. The plant was then identified and authenticated by a botanist and the sample specimen deposited in Herbarium unit of Department of Biology, College of Computational and Natural Science, University of Gondar for future reference with voucher number of DG0024/210.
3.1.3. Experimental AnimalsSwiss albino mice of either sex 6-8 weeks (25–35 g) from Pharmacology and Toxicology laboratory, School of Pharmacy, College of Health Sciences, Mekelle University were used in the present study. Experimental animals were housed in polypropylene cage and maintained under standard animal house condition (at ambient temperature, and with a 12 h light-dark cycle) and allowed free access to the standard pellet diet and water ad libitum. Before initiation of the experiment the animals were acclimatized to laboratory condition for a period of seven days.
All procedures have been undertaken as per the guide for the care and use of laboratory animals (OECD, 2008). The study protocol was approved by Health Research Ethics Review Committee (HRERC), College of Health Sciences, Mekelle University with protocol number (ERC1206/2018). Animals were scarified under anesthesia after completion of the experiments. 3.2.
Methods3.2.1. Extraction of Plant MaterialThe roots were cleaned from the dust and debris and washed gently with water; reduced to appropriate size and air dried under shade for two weeks.
The dried root material was then pulverized with grinder. Figure 4: The extraction process of Cucumis ficifolius roots using 80% methanol.Then a total amount 1.335 kg of dried root powder was macerated in methanol (80%) for 72 hours (Bantie et al., 2014). The extraction process was facilitated by using an orbital shaker at 120 rpm. The mixture was first filtered using muslin cloth and then with Whatman filter paper no.1.
Re-maceration of the remaining residue was done for another 72 hours twice to obtain maximal yield. The filtrates were concentrated in oven dryer set at 40 oC. The dried powder of the extract was kept in a refrigerator at 4 oC in air-tight containers until use.
3.2.2. Solvent-Solvent ExtractionThe procedure of solvent-solvent fraction was carried out using a separatory funnel. A total of 45 g of the crude extract was dissolved in 270 ml of distilled water. Then extraction was performed successively using solvents of increasing polarity, starting from chloroform (270 ml X 3) then n-butanol (270 ml X 3).
After collecting the chloroform and n-butanol fractions the remaining residue was considered as aqueous fraction. The fractions were concentrated using oven dryer at 40°C. The dried powders were kept in air-tight containers wrapped with the aluminum foil and stored in a refrigerator at 4 oC until further use. 3.2.3 Acute Oral Toxicity Test The acute oral toxicity test was carried out according to limit test recommendations of Organization for Economic Co-operation and Development (OECD) 425 Guideline (OECD, 2008).
Nulliparous, non-pregnant and healthy female Swiss albino mice (age of 8-12 weeks) weighing from 20-30gm were employed for this test. A total of five female animals were used. The extract to be tested was calculated based on fasting body weight and volume administered was determined based on OECD guideline that states 1 mL/100 g of body weight of the animal. On day one, a mouse fasted for four hours was orally given 2000 mg/kg of the extract dissolved in distilled water by oral gavage. The mouse was then observed for physical or behavioral changes at least once during the first 30 min, periodically for 24 h, with special attention during the first 4 h.
After 24 h other four mice were selected and fasted for 4 h and administered a single dose of 2000 mg/kg. The mouse was then observed for at least once during the first 30 min, periodically for 24 h, with special attention during the first 4 h, and daily thereafter for a total of 14 days Animals were observed for any manifestations including tremors, convulsions, salivation, diarrhea, lethargy, sleep and coma.3.2.4 Anti-nociceptive Activity Testi. Acetic-Acid Induced Writhing TestThe test was carried out as per the method described by Birhane et al., (2015) with slight modification.
Mice of either sex (26-35 gm) were used. Animals fasted for 12 hours were randomly divided into five and six groups of six mice each for evaluation of the crude extract and fractions, respectively. The negative control group was administered with distilled water (DW) 10 ml/kg, whereas the positive control group received 150 mg/kg aspirin.
The remaining 3 groups were treated with the crude extract at doses of 200, 400, and 800 mg/kg. The 200, 400 and 800 mg/kg doses were selected based on the acute toxicity study. In fractions, six groups of animals were used: Group I received DW (10ml/kg); Group II and III were treated with 100 mg/kg and 200 mg/kg aqueous fraction, respectively; Group IV was administered with 100 mg/kg butanol fraction and Group V (200 mg/kg, butanol fraction) while Group VI was treated with aspirin 150 mg/kg. Thirty minutes following administration of vehicle, standard drug and test substance; the animals were subjected to intraperitoneal (i.p) injection of acetic acid solution (0.
6%, 10 ml/kg) (Debebe et al., 2007). The writhing response which consists of a contraction of the abdominal muscle together with a stretching of the hind limbs and arching of the back were used as a writhing response and measured for 20 min after a latency period of 5 min.
Percent protection was calculated by applying the formula (Tadiwos et al., 2017).% Analgesic activity=(Mean no.writhes (control) -Mean no.of writhes (treated))/(Mean no.
of writhes (control)) ×100ii. Hot Plate TestThe hot plate test was performed according to the method described by Debebe et al. (2007) with a slight modification using hot-plate apparatus (ORCHID Scientific, India) which was maintained at 55 ± 0.1 °C. Overnight fasted animals were randomly assigned into different groups.
The positive control group received morphine 20 mg/kg orally. Details of grouping and dosing of animals is presented in Table 1. The baseline latencies were determined twice at 15 minute intervals and the first readings were discarded. Latencies were then determined at 30, 60, 90 and 120 minutes after test substance, vehicle or standard drug administration. A cut off time of 20 secs was considered by taking three times the mean pre-drug latency was imposed to minimize tissue damage (Debebe et al., 2007). The nociceptive latency in seconds was quantified by considering the interval between the instant the animal reached the hot plate and licked its paw or jumping off the hot plate.
The post-drug latency: T1 was estimated according to the reaction time of each mouse at 30, 60, 90 and 120 minutes after treatment. T0 represented the mean pre-drug latencies. For each group, the percentage of protection against thermal stimulus was determined by using the formula described by (Yonathan et al.
, 2006).% Protection against thermal stimulus=(T0-T1)/T0 ×100iii. Formalin Test Formalin test was performed using the method developed by and Hunskaar et al, 1986. Mice of either sexes fasted overnight with the provision of water were used. Mice of either sex were fasted overnight and randomly selected and assigned into groups of five each with six mice. Group I received DW (10 ml/kg) was served as the negative control while group II received aspirin at dose of 200 mg/kg served as positive control. The remaining groups (III to V) were given the test extract at a dose of 200, 400 and 800 mg/kg. Grouping and dosing used in solvent fractions is also indicated in (Table 3).
Mice in each group was allowed for at least 20 minutes acclimatization in transparent observation cage before starting the experiment (Chung et al., 2000). Then 0.02 mL of 5 % formalin solution was administered by intraplantar injection in to the mice dorsal surface of the right hind paw. Two phases of nociception namely, the early and late phase were observed during the course of the experiment. The first phase was recorded by taking the time of the animal spent licking the paw for 0-5 minutes after injection of formalin (Hunskaar et al.
, 1985) while the second phase was recorded by taking the time of the animal spent licking the paw for 15-30 min after formalin injection (Santos et al., 1998; Adedapo et al., 2014). The percentage of inhibition nociception for the two phases was calculated using the following formula (Chung et al., 2000) % Inhibition=(Control mean-Test mean)/(Control mean) ×100 Table 1: Grouping and dosing of mice used on formalin testGroup Dose (ml/kg or mg/kg) Group Dose (ml/kg or mg/kg)DW 10 DW 10Crude methanolic extract 200 Aqueous fraction 100 400 200 800 Butanol fraction 100Aspirin 200 200 Aspirin 200 3.2.5 Evaluation of Anti-inflammatory Activity Anti-inflammatory activity of the 80% methanolic crude root extract and solvent fractions of C.
ficifolius was performed using the method described by Winter et al. (1962) with slight modification. Swiss albino mice of either sex weighting from 20-37gm were used. Animals fasted overnight were randomly assigned into nine groups each with six mice Swiss albino mice of either sex (27-37gm) were used (Table 2). Thirty minutes before injection of carrageenan, distilled water (DW, 10 mL/kg), the test substances and aspirin (200mg/kg) were administered orally. Just before induction of inflammation the leg of each mouse was marked on the skin over the lateral maleous (Winter et al.
, 1962). The basal volume of the right hind paw of individual mouse was measured with digital plethysmometer (PLM 02). Then 0.05 mL of 1% carrageenan in normal saline was injected in to the dorsal surface of the right hind paw. The volume of injected paw was measured at 1, 2, 3 and 4 hours after carrageenan injection.Paw diameter before carrageenan injection was compared with the same paw diameter after administration of carrageenan by calculating the percentage inhibition applying the following formula (Masresha et al., 2012). % inflammation inhibition =((Vt-Vo) control-(Vt-Vo) treated )/((Vt-Vo) control) ×100Where, Vt =the mean paw volume in control and drug treated group at time t Vo =the mean paw volume in control and treated group at time 0 In addition the increase in paw volume, i.
e., inflammation expressed in percentage was calculated according to the formula given by (Masresha et al., 2012).%inflammation (%I)=(Vf-Vi)/( Vi) ×100 Where, Vi is the mean volume of the paw before carrageenan injection and Vf, the volume of the paw after carrageenan injection.Table 2: Grouping and dosing of mice used anti-inflammatory activities of crude extract and solvent fractions of Cucumis ficifolius.
Groups Treatments Dose (mg/kg or ml/kg)GI Distilled water 10 GII Methanol extract 200GIII Methanol extract 400GIV Methanol extract 800GV Aqueous fraction 100GVI Aqueous fraction 200GVII Butanol fraction 100GVIII Butanol fraction 200GIX Aspirin 2003.2.6 Statistical AnalysisData was entered and analyzed with the IBM statistical package for social sciences (SPSS) version 21.
The data obtained in the study was tabulated and expressed as mean ± standard errors of the mean (SEM). Then statistical analysis was carried out using one-way analysis of variance (ANOVA) followed by Tukey post-hoc test to compare variations among groups. The result was considered significant when p < 0.05.