Lab 2 Immunocytochemistry

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University of Kentucky *

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210

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Biology

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Feb 20, 2024

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docx

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BMB 210 Lab 2 LABORATORY 2 IMMUNOCYTOCHEMISTRY - PART 2 Before lecture, start the rinsing procedure as described on page 2, # 1&2. In today’s lab, you will complete the immunocytochemistry experiment that you began last week. INTRODUCTION Recall that in last week’s lab, you began the incubation of rat pancreas sections with rabbit anti-insulin and mouse anti-glucagon primary antibodies. Today, after rinsing the sections in KPBS to remove unbound primary antibodies, you will add the secondary antibody solution. Two different secondary antibodies will be used: donkey anti-rabbit and donkey anti-mouse. These antibodies were generated by injecting donkeys with either rabbit or mouse antibody proteins. The donkey immune system produced antibodies against the foreign proteins, and these donkey antibodies will be used as the secondary antibodies that bind to the primary antibodies . Several secondary antibodies can bind to each primary antibody (see Fig. 1). Antibodies are not visible through the microscope unless they are tagged with some type of marker. Covalently attached to the secondary antibodies are fluorescent molecules that can be seen using a fluorescence microscope. These fluorescent molecules contain electrons that are excited by certain wavelengths of light. As the excited electrons return to their ground state, photons of a specific wavelength are emitted and are seen as a particular color. The fluorescence microscope contains filters that allow the passage of certain wavelengths and not others. The donkey anti-rabbit secondary antibodies are tagged with A488, which fluoresces green. The donkey anti-mouse secondary antibodies are linked to Rhodamine Red-X, which fluoresces red. Cells producing insulin or glucagon, therefore, can be seen through the microscope via a series of antibodies and fluorescent markers. What color should insulin- producing vs. glucagon-producing cells appear? Secondary antibodies (donkey anti-rabbit or anti-mouse) with fluorescent markers attached Primary antibody (rabbit anti-insulin or mouse anti- glucagon) Antigen molecule in tissue (insulin or glucagon) Fig.1. Diagram illustrating the consecutive binding of primary and secondary antibodies to an antigen in a tissue. Page 1 of 3
BMB 210 Lab 2 EXPERIMENTAL PROCEDURE – Wear Gloves! Secondary Antibody Incubation 1. Remove the slides from the humid chamber one at a time. Turn the slide on end to allow the puddle of primary antibody to run onto a paper towel, and place the slide in a slide carrier inside a staining boat filled with 1X KPBS. Do not let the slides dry. 2. Rinse the slides in 1X KPBS two times for 5 minutes each rinse (2 X 5). After the final slide is put into KPBS from step 1, begin timing. This is rinse #1. After 5 minutes, gently remove the slide carrier and pour the KPBS into the sink. Pour fresh 1X KPBS into the boat and return the slide carrier to the boat. This is rinse #2. 3. After the second 5 minute rinse, remove the slides one at a time, wipe the excess KPBS from the back with a Kimwipe as you did last week, and set the slide in the humid chamber. Add 100 l of secondary antibody solution onto the tissue inside the hydrophobic/PAP ring. Repeat for all slides. DO NOT LET THE TISSUES DRY! The secondary antibody solution contains donkey anti-rabbit linked to a green marker, A488 (A488 D R) and donkey anti-mouse linked to Rhodamine Red-X (RedX D Ms). The secondary antibody was diluted in the same dilution buffer used last week. 4. Cover the humid chamber and let the slides incubate at room temperature for at least 1 hour. ***During the 1-hour incubation, we will complete the primary research article citation and reference formatting activity.*** Cover slipping and observation under the microscope After the slides have incubated for one hour: 5. Remove one slide at a time from the humid chamber, and blot the secondary antibody solution onto a paper towel as you did above. Rinse the slides in 1X KPBS, 2 X 5, as described in steps #1 and 2 above. 6. Coverslip with an aqueous mounting medium: After the last rinse, remove one slide at a time and lay it on a paper towel -- specimen side up! Using a Pasteur pipette, add 2 drops of mounting medium (1:1 KPBS/glycerol) over the tissue on the slide. Coverslip with a 24 X 40 mm coverslip, avoiding air bubbles as much as possible. 7. Lay the slides horizontally in a plastic slide tray. When handling the slides, always hold them horizontally or the coverslip will slide off. Your instructor will take you to the fluorescence microscope in the “imaging room” in Y103 to look at your slides. Page 2 of 3
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