MOMA-2 IHC Staining | Lusis Laboratory

MOMA-2 IHC Staining for Macrophages in Aortic Sections (ABC Method)

MATERIALS

Pap pen
Coplin Jars
Dry Acetone
1x PBS
BSA (Sigma Cat. No. A-7030)
Normal Rabbit Serum (Vector S-5000)
MOMA-2 Primary Antibody (Accurate Chemical and Scientific Corporation Cat. No. BMAMOMA2)
Biotinylated rabbit Anti-rat IgG monoclonal Ab (Vector, Cat. No. BA 4001, 0.5mg, in liquid form)
ABC-AP Reagent - Alkaline Phosphatase Complex (Vector AK – 5000)
Vector Red Substance - Substrate for Alkaline Phosphatase in ABC-AP complex (Vector SK-5100)
Levamisole Solution (Vector SP - 5000)
Aqueous Hematoxylin (Biomeda M10)
Sodium Bicarbonate (Fisher S233-500, 500mg)
Fast Green Dye (optional, can get small aliquot from Xiuping)
Glycerol Mount (Sigma Cat. No. GG1-15ML)

PROTOCOL

Bring Slides to room temperature for 30 minutes to 1 hour.
Circle Sections with Pap Pen (Research products, hydrophobic slide marker) - Circle each section (3 sections per slide) around with pap pen, making sure that the boundaries of neighboring sections do not overlap. Press lightly as you mark but mark well, going around each circle several times.

Note: Make sure that there is no overlap into neighbor sections and that all are well circled and sealed, otherwise, when you add components, everything will be mixed between sections.

Fix Slides - Submerge slides in chamber filled with Dry Acetone for 15 min at Room Temperature.
PBS Wash - 3 washes in PBS (prepared fresh), 10 minutes each.
- Remove slides from acetone and tap a little bit on dry tissue to remove excess acetone.
- Submerge immediately in fresh PBS. Transfer should be quick so that sections do not dry.
- After each wash, change to fresh PBS. Two washes are enough.
Block - Blocking should be performed using serum from the species of the 2° Ab. Since the secondary antibody is less specific, it may bind to certain regions of cells resembling its specificity. Blocking this with serum from the same species should reduce this effect.
- 3% BSA - 1.2g BSA in 40ml PBS
- 5% Blocking Solution - 0.5ml Normal Rabbit Serum (NRS) in 10ml 3%BSA Solution (a 20x dilution in 3%BSA). Mix by gently inverting, keep on ice. This solution should be made fresh each time
- After PBS wash, place slides on moisture chamber, not directly touching moist surface. Tap excess PBS off on tissue paper.
- Dry surface of slides using a cotton swab. Make sure regions around slides are perfectly dry, going around hydrophobic mark. If necessary, re-mark sections using pap pen. Do not touch sections with swabs.
- Add about 150μl drops Blocking Solution (approx 3 drops if using a Pasteur pipet). Add solution slowly, making a bubble, without touching the sections with pipet. Make sure solution does not spill into neighboring sections.
- Cover box and incubate for 2 hours in the dark.
Primary Antibody - MOMA-2 at 1:400 Dilution
- Rinse Wash Buffer - Make this buffer first by adding 1 part Blocking Solution: 4 parts PBS, (or a 5x dilution in PBS): Ex: 1ml BS + 4ml PBS, make fresh each time.
- Calculate amount of Ab solution you will need. Need about 150μl per section (Ex: 150μl/section X 3sections/slide = 450μl/slide;
  450μl/slide X 6 slides = 2.7ml; Make about 3ml
- Make 400x dilution of MOMA-2 antibody in the Rinse Wash Buffer
- V₂/V₁ = DF; 3ml/V₁ = 400X; V₁ = 3ml/400 = 7.5μl AB
- Dilute 7.5μl of MOMA-2 Ab in 3ml of Rinse Wash Buffer. Mix by inversion and place on ice.
- Remove Blocking solution from each section, by tapping off. Dry around slides carefully; re-draw with pap pen if necessary.
- Add about 150µl of 1° Ab to each section, without leaking or touching sections. If leaky, dry with cotton swab and re-draw hydrophobic mark.
- Control - Add RW buffer, instead of primary antibody to one section. No staining should be seen in this section.
- Cover and incubate at RT for 1 hour.
PBS Wash - 2X, 10 minutes each, 1X with Rinse Wash buffer for 10 minutes. Add RW with pipet.
Secondary Antibody - Biotinylated rabbit Anti-rat IgG monoclonal Ab (Vector, Cat. No. BA 4001, 0.5mg, in liquid form)
- Make a 200X Dilution of secondary antibody in RW buffer. Need about 3ml.
- V₂/V₁ = 200X; V₁ = 3ml/200 = 15μl 2° Ab
- Dilute 15μl of secondary antibody in 3ml RW buffer
- After PBS wash, place slides in moisture chamber. Dry around sections. Add 150μl (2 drops) of RW buffer to each (previous step). Remove RW by tapping. If you have many slides, do this in parts so they do not dry (i.e. take out a few, dry and add RW, then take out some more).
- Add 150μl of 2° Ab (2-3 drops) in a bubble so they don’t leak to other sections. Cover and incubate in the dark for 1 hour
  RT
PBS Wash - 3X, 5 minutes each, RT.
ABC-AP Reagent
- Calculate amount you will need:
- 5ml PBS, 1 drops A, 1 drops B. Mix in bottle of kit and cap, shake bottle and place on ice. Incubate for at least 30min in the dark before using. Make fresh each time.
- After PBS wash shake off excess by tapping on napkin, dry back with Kim wipe and place on moist chamber. Dry around sections.
- Add 2 drops (2 is enough, can go back and add more if you have left) of ABC-AP reagent to each section.
- Cover and incubate in moist chamber 1 hour RT in dark
PBS Wash - 2X PBS, 5 minutes each. 1X ddH₂O.
Vector Red Substance - Substrate for AP in ABC-AP complex:
- 5ml Tris-HCl Buffer (pH 8.3)
- 1 drops Levamisole Solution (Vector SP – 5000)
- 2 drops each of 1, 2, 3
If 5ml make in plastic tube provided, if more, in 15ml tube. Mix by inversion and gentle vortex. Mix and keep in the dark
- Remove slides from moist box, shake off excess and dry around sections. Place in a white surface so color development if easier to see.
- Add 2 drops each section. Incubate with reagent in dark, RT
- Check for development until you start seeing a reddish color
- If 10μm sections, and you start seeing color development really fast, leave for 5-10 minutes, maybe less. Need to check constantly, under microscope, so you note if background-staining starts to develop also. If you start seeing background, stop developing.
- If you remove too early, color will not be strong enough for contrast. If too late, you will start seeing background. Need to constantly check development.
- Control should have no color.
Counter stain
- When ready, shake off excess reagent, tapping on napkin.
- Dip sections in ddH₂O; it does not matter how long.
- Dip section in Hematoxylin, which will stain all nuclei purple. If you have many slides, use chamber, if few, add 2 drops each section. If used Hematoxylin, make sure you filter before use. Incubate 4-6 min at RT, If new, 2 min would be ok. (Aqueous Hematoxylin, Biomeda M10)
- Rinse with a lot of tap water, dipping slides in large container with water and rinsing.
- Dip slides in Saturated Sodium Bicarbonate for 1 minute. This will turn counterstaining to a bluish tone, so that the contrast between red (macrophages) and blue (all else) will be better. Easier to see than red:purple contrast. (Sodium Bicarbonate, Fisher S233-500, 500mg).
- Rinse with tap water again.
- Fast green - Optional staining. Filter Fast green before use. Dip in fast green for 3 minutes. Then rinse profusely in tap water.
Mounting: Glycerol Mount
- Let slides dry at RT, then place on hot bed at ~40°C to warm
- Warm mounting media to 55°C on a beaker over a hot plate; media should become liquid. Keep mounting media there until ready to use
- Invert mounting media to let bubbles go up, take first drop in Kim wipe
- Add one drop of Glycerol mount to each section. Carefully, not to touch sections, and without bubbles. If the slide and the mounting media are warm as they should be, the media will spread out on its own
- To place coverslip, place edge of it on right side of slide and push down slowly with forceps. Go slowly, pushing down toward left edge of slide, this will allow bubbles to get out
- Make sure all sections are covered with mounting media, push out any bubbles remaining
- Dry overnight, not tilting slide or mounting media may drip.