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S. S. College, Jehanabad Department: Zoology Class: M.Sc S. S. College, Jehanabad Department: Zoology Class: M.Sc. Semester II Subject: Zoology Topic: Differentiation of neutral lipid by Nile blue sulfate method Mode of teaching: Google classroom & WhatsApp Date & Time: 06.08.2020 & 10:30 Teacher: Praveen Deepak To join Department’s group, students can use following link https://chat.whatsapp.com/EHuHNfQzoAzJBMFNJvsjQx or scan QR Code WhatsApp No.: +91 75360 68068 DIFFERENTIATION OF NEUTRAL LIPID BY NILE BLUE SULFATE METHOD ____________________________________________________________ Nile blue (hydrogen sulfate) belongs to the group of oxazine dyes. Nile blue (hydrogen sulfate) is used as a single dye to differentiate neutral fats and cholesterol esters from free fatty acids and phospholipids. It was first introduced by Lorrain Smith in 1908 for distinguishing neutral lipids (triglycerides) from fatty acids. However, later Kaufmann and Lehmann in 1926 concluded that it was of no histochemical value after applying it to a number of lipois. Lison in 1935 concluded that the red colouration is characteristic of lipoids in general after a profound histochemical study. Now, it is utilized in the differentiating natural lipids from fatty acids because neutral lipids stain red, while fatty acids stain dark blue with the nile blue sulfate. This test ensures that if a substance is known beforehand to be lipoid and colours red with nile blue, it consists of neutral lipoids (esters and/or hydrocarbons); if it colours blue, it may contain these, but acidic lipoids (fatty acids, phospholipines, and perhaps some others) are certainly present as well. The presence or absence of cholesterol cannot be established with nile blue. Principle Nile blue is the composition of two dyes that is blue oxazine and red oxazone (an oxidation product of oxazine). Oxazone is a lysochrome i.e. fat soluble dyes, which reacts with the neutral lipids to give a red to pink colour. For phospholipids, calcium compounds of phospholipid combine with the oxazine form of Nile blue sulfate and survive subsequent treatment, neutral lipids are dissolved out by acetone, and proteins and other interfering substances are destained by the acetic acid and hydrochloric acid baths. Fixation and Tissue Processing Tissue sections of 3 – 5 µm thickness are formalin fixed (formalin containing 1% CaCl2) and processed for paraffin infiltration and embedding or freeze formalin (containing 1% CaCl2 as in the case of preparation for paraffin embedding) fixed or 8µm thick tissue sections are freeze fixed and proceeded for mounting with suitable aqueous mounting media. Freeze formalin fixation is opted and done with following steps: • Specimen is placed in tissue cassette, wash in running water for 5 minutes. • Tissue is removed tissue cassette; blot well, removing all excess water from tissue. • Tissue is frozen according to tissue freezing manual. Equipment • Slides • 22mm square coverslips • 250 ml beaker • 250 ml volumetric flask • Whatman #4 filter paper • Ceramic staining rack • Staining dish • Coplin staining jar Praveen Deepak, Department of Zoology, S. S. Colleg e, Jehanabad • Forceps • Latex globes Reagents • Nile blue • Anhydrous Calcium chloride (CaCl2) • Sulfuric acid (H2SO4) • Glacial acetic acid • Conc. hydrochloric acid (HCl) • Acetone • Gelatin or carbowax • Glycerine • Phenol • Absolute alcohol • Distilled water Solution 1. Nile blue solution • Prepared by adding 0.5gm of Nile blue in 99ml of distilled water and mix well. • Finally, resulting solution is added with 1ml of 0.5% sulfuric acid. 2. HCl solution (0.5%) • Prepared by adding slowly 21ml of conc. HCl in 400ml distilled water. • Further distilled water is added to adjust the volume upto 500ml. 3. Calcium chloride solution (1%) • Prepared by dissolving 2.9gm of anhydrous calcium chloride (CaCl2) in a mixture of 95ml of distilled water and 5ml of dilute hydrochloric acid. • Osmolarity is measured using an osmometer; if below 100mmol/kg, more CaCl2 is added, and if above 1000, more distilled water is added. 4. Acetic acid solution (5%) • Prepared by adding 5ml glacial acetic acid into 95ml of distilled water and mix thoroughly. • Unused acetic acid is being discarded at the end of the day. 5. Glycerin gelly • Prepared by mixing 10gm of gelatin in 60ml of distilled water with gentle heating. 2 | P a g e • After dissolving of gelatin completely, 1.0ml of phenol is added in 70ml of glycerin. Procedure 1. Fix 6–12 hr. in 10% formalin containing 1% CaCl2. 2. Cut frozen sections without embedding or after gelatin or carbowax. 3. Stain 90 minutes at 60°C in saturated aqueous Nile blue sulfate solution. 4. Rinse in distilled water, and place in acetone heated to 50°C. 5. Remove the acetone from the source of heat and allow the sections to remain 30 min. 6. Differentiate in 5% acetic acid 30 min. 7. Rinse in distilled water, and refine the differentiation in 0.5% HCl for 3 min. 8. Wash in several changes of distilled water. 9. Mount in glycerol jelly. Results Neutral lipids appeared to be stained with dark red to pink colour and acidic lipids such as phospholipids found to be stained with blue in colour. Other areas appeared to be unstained under microscopic observation. References 1. Menschik Z. Nile blue histochemical method for phospholipids. Biotechnik Histochemistry. 1953; 28(1): 13– 8. 3 | P a g e 2. Cain A. J. The use of Nile blue in the examination of lipoids. Q. J. M. S. 88(3): 383–392. 3. Singh A. Histochemical techniques to demonstrate lipids. https://conductscience.com/histochemi cal-techniques-to-demonstrate-lipids/ ∞∞∞∞∞ 4 | P a g e .
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