You've got your box of Ahelixbio pre-cast gels. You've checked the compatibility. You're ready to run your Western blot.
But now what?
How do you open the pouch? Do you need to rinse the wells? What voltage should you use? How long will it take?
Don't worry. Using pre-cast gels is much easier than hand-casting—but there are a few critical steps you can't skip.
In this guide, I'll walk you through the complete step-by-step protocol for using Ahelixbio pre-cast gels, from opening the pouch to transferring your proteins for Western blot.
What You'll Need
Before You Start: Preparation
Step 1: Remove Gel from Packaging
Step 2: Set Up the Electrophoresis Tank
Step 3: Prepare Your Samples
Step 4: Load the Gel
Step 5: Run the Gel
Step 6: Remove the Gel from Cassette
Step 7: Transfer for Western Blot
Step 8: Clean Up and Storage
Quick Protocol Card
Frequently Asked Questions
| Item | Notes |
|---|---|
| Ahelixbio pre-cast gel | Appropriate percentage for your protein |
| Electrophoresis tank | Compatible with Ahelixbio gels (e.g., Bio-Rad Mini-PROTEAN) |
| Power supply | Capable of 80–200V |
| Pipettes and gel-loading tips | Long, thin tips for wells |
| Syringe (1 mL) with blunt needle | For rinsing wells |
| Spatula or gel knife | To open cassette |
| Rocking platform | For staining/blocking |
| Transfer apparatus | For Western blot |
| Reagent | Notes |
|---|---|
| 1X Running buffer | Tris-Glycine or compatible |
| Protein samples | In loading buffer, boiled |
| Protein ladder | Pre-stained recommended |
| 1X Transfer buffer | For Western blot |
| Membrane (PVDF or nitrocellulose) | For Western blot |
Samples mixed with loading buffer (1:1 or as appropriate)
Reducing agent added (β-ME or DTT) – fresh is best
Boiled at 95°C for 5–10 minutes
Centrifuged (if debris present)
Kept on ice until loading
Before opening the pouch, check:
| Check | What to Look For | Action if Bad |
|---|---|---|
| Expiration date | At least 3 months remaining | Use soon or return |
| Pouch integrity | No punctures or tears | Don't use |
| Storage temperature | Gel feels cold (4°C) | If warm, check quality |
| Visible damage | Cracks, cloudiness through pouch | Don't use |
| Step | Instructions |
|---|---|
| 1 | Remove gel pouch from 4°C refrigerator |
| 2 | Let sit at room temperature for 5–10 minutes |
| 3 | Do NOT open pouch during warming (prevents condensation) |
| Buffer Type | Recipe (1X) | Notes |
|---|---|---|
| Tris-Glycine | 25 mM Tris, 192 mM glycine, 0.1% SDS | Most common |
| Tris-Tricine | For small proteins (<20 kDa) | Use with appropriate gels |
| MOPS/MES | For Bis-Tris systems | Check compatibility |
Pro tip: Make fresh 1X running buffer for each run. Reusing buffer causes smearing and slow migration.
| Step | Instructions | Image (Description) |
|---|---|---|
| 1 | Locate the notch or tear line on the pouch | Top edge of foil pouch |
| 2 | Tear open carefully (or use scissors) | Avoid touching the gel |
| 3 | Remove the gel cassette | Hold by edges only |
| Step | Instructions |
|---|---|
| 1 | Hold the cassette firmly by the sides |
| 2 | Grasp the comb at both ends (not middle) |
| 3 | Pull slowly and evenly straight up |
| 4 | Do NOT wiggle or pull at an angle |
| Step | Instructions |
|---|---|
| 1 | Locate the green or white tape at the bottom of the cassette |
| 2 | Peel off slowly and evenly |
| 3 | Check that the tape comes off cleanly (no residue) |
This is the most important step that beginners skip.
| Step | Instructions |
|---|---|
| 1 | Fill a 1 mL syringe with 1X running buffer |
| 2 | Attach a blunt needle or gel-loading tip |
| 3 | Insert tip into the first well, all the way to the bottom |
| 4 | Gently flush buffer through the well |
| 5 | Repeat 2–3 times per well |
| 6 | Repeat for all wells |
Why this matters: Rinsing removes:
Unpolymerized acrylamide (can cause smearing)
Air bubbles (prevent sample from settling)
Storage buffer residues
Warning: Skipping the rinse step is the #1 cause of bubble problems and smearing with pre-cast gels.
| Step | Instructions |
|---|---|
| 1 | Place the gel cassette into the tank with the short plate facing inward |
| 2 | Push down gently until the cassette is fully seated |
| 3 | If running two gels, insert second cassette on the other side |
| Step | Instructions |
|---|---|
| 1 | Pour 1X running buffer into the inner chamber |
| 2 | Fill until buffer covers the wells completely |
| 3 | Check for leaks (wait 30 seconds) |
| Step | Instructions |
|---|---|
| 1 | Pour 1X running buffer into the outer chamber |
| 2 | Fill to the level indicated on your tank (usually between the "min" and "max" lines) |
| 3 | Ensure the bottom of the gel is submerged |
| Step | Instructions |
|---|---|
| 1 | Wait 2 minutes after filling inner chamber |
| 2 | Check if buffer level has dropped |
| 3 | If leaking, reseat the gel and try again |
| Step | Instructions | Time |
|---|---|---|
| 1 | Thaw samples on ice | 5–10 min |
| 2 | Mix sample with 2X or 4X loading buffer (1:1 or 3:1 ratio) | 1 min |
| 3 | Add fresh reducing agent (β-ME or DTT) if needed | 1 min |
| 4 | Boil at 95°C for 5–10 minutes | 5–10 min |
| 5 | Centrifuge at 14,000 rpm for 5 minutes (if debris present) | 5 min |
| 6 | Keep on ice until loading | – |
| Component | Amount for 10 mL |
|---|---|
| Tris-HCl (pH 6.8) | 2.5 mL of 0.5 M |
| SDS | 0.8 g |
| Glycerol | 4 mL |
| β-mercaptoethanol | 0.4 mL |
| Bromophenol blue | 0.002 g |
| dH₂O | to 10 mL |
| Gel Thickness | Well Format | Max Volume | Recommended Volume |
|---|---|---|---|
| 1.0 mm | 10-well | 30 µL | 20–25 µL |
| 1.0 mm | 12-well | 25 µL | 15–20 µL |
| 1.0 mm | 15-well | 20 µL | 10–15 µL |
| 1.5 mm | 10-well | 60 µL | 40–50 µL |
| 1.5 mm | 12-well | 50 µL | 30–40 µL |
| 1.5 mm | 15-well | 40 µL | 20–30 µL |
| Sample Type | Recommended Protein per Lane |
|---|---|
| Cell lysate (routine Western) | 20–30 µg |
| Cell lysate (weak antibody) | 40–50 µg |
| Purified protein | 0.5–5 µg |
| Tissue lysate | 30–50 µg |
| Step | Instructions |
|---|---|
| 1 | Use gel-loading tips (long, thin) |
| 2 | Draw sample into tip (slightly less than well volume) |
| 3 | Insert tip just below the top edge of the well |
| 4 | Expel sample slowly – do not squirt |
| 5 | Watch the sample settle to the bottom |
| 6 | Remove tip slowly |
| 7 | Move to next well |
| Lane | Sample | Volume |
|---|---|---|
| 1 | Protein ladder | 5–10 µL |
| 2 | Empty or loading buffer | – |
| 3 | Sample 1 | As needed |
| 4 | Sample 2 | As needed |
| 5 | Sample 3 | As needed |
| 6 | Sample 4 | As needed |
| 7 | Sample 5 | As needed |
| 8 | Sample 6 | As needed |
| 9 | Empty or loading buffer | – |
| 10 | Protein ladder (optional) | 5–10 µL |
Pro tip: Load ladder in the first and last lanes. This helps identify "smiling" (curved bands) if it occurs.
| If you have empty lanes... | Do this |
|---|---|
| 1–2 empty lanes | Load 1X loading buffer (same volume as samples) |
| 3+ empty lanes | Consider re-arranging samples to minimize empty lanes |
| Why? | Prevents "edge effects" (distorted bands on ends) |
| Phase | Voltage | Time | Stop When |
|---|---|---|---|
| Stacking (optional) | 80–100V | ~20 min | Dye front enters resolving gel |
| Resolving | 120–150V | 30–60 min | Dye front reaches bottom |
Simpler approach: Run at 120V constant for the entire run. This works well for most applications.
| Gel Type | Voltage | Approximate Time |
|---|---|---|
| 4–20% gradient | 120–150V | 45–60 min |
| 10% fixed | 120–150V | 40–50 min |
| 12% fixed | 120–150V | 45–55 min |
| 15% fixed | 120–150V | 50–65 min |
| What to Watch | What It Means |
|---|---|
| Bubbles rising from wells | Current is flowing (good) |
| Dye front (blue line) | Tracking migration progress |
| Temperature of tank | Should be warm, not hot |
| Buffer level | Should not drop significantly |
| Stop when... | Why |
|---|---|
| Dye front reaches the bottom of the gel | Optimal separation |
| Your smallest protein of interest has separated | Don't let it run off |
| You see smiling bands starting | Stop before they worsen |
Never let the dye front run off the gel if you plan to stain with Coomassie – it carries small proteins with it.
| Step | Instructions |
|---|---|
| 1 | Turn off power supply and disconnect leads |
| 2 | Remove the tank lid |
| 3 | Pour out running buffer (discard) |
| 4 | Remove gel cassette(s) from tank |
| Step | Instructions | Caution |
|---|---|---|
| 1 | Place cassette on a clean, flat surface | Short plate facing up |
| 2 | Insert a spatula or gel knife into the corner | Between the two plates |
| 3 | Gently twist to separate the plates | Don't force |
| 4 | Run the spatula along both sides | To fully separate |
| 5 | Lift off the short plate | Gel will stick to long plate |
| Step | Instructions |
|---|---|
| 1 | Locate the interface between stacking and resolving gel |
| 2 | Use a clean razor blade or spatula |
| 3 | Cut straight across just above the resolving gel |
| 4 | Discard the stacking gel (contains no target proteins) |
| Step | Instructions |
|---|---|
| 1 | Cut a small corner of the gel (for orientation) |
| 2 | Carefully transfer to staining tray or transfer apparatus |
| 3 | Keep gel moist with transfer buffer or water |
| Step | Instructions |
|---|---|
| 1 | Equilibrate gel in 1X transfer buffer for 5–10 min |
| 2 | Activate PVDF membrane in methanol (15 sec) if using PVDF |
| 3 | Equilibrate membrane and filter paper in transfer buffer |
| 4 | Assemble sandwich: Filter paper → Gel → Membrane → Filter paper |
| 5 | Roll out all bubbles with a pipette or roller |
| 6 | Place sandwich in transfer tank |
| 7 | Fill with cold transfer buffer |
| 8 | Transfer at 100V for 60 min (or 30V overnight at 4°C) |
| Protein Size | Transfer Method | Time | Methanol % |
|---|---|---|---|
| <20 kDa | Wet (100V) | 30–45 min | 20% |
| 20–80 kDa | Wet (100V) | 60 min | 20% |
| 80–150 kDa | Wet (100V) | 90 min | 10–20% |
| >150 kDa | Wet (30V overnight) | 12–16 hours | 10% |
| Step | Instructions |
|---|---|
| 1 | After transfer, place membrane in Ponceau S solution |
| 2 | Stain for 2 minutes |
| 3 | Rinse with water |
| 4 | Check for protein bands (should see ladder and sample bands) |
| 5 | If no bands → transfer failed |
| 6 | If bands present → proceed to blocking |
| Item | Disposal Method |
|---|---|
| Used gel | Solid waste (polymerized gel is non-toxic) |
| Running buffer | Down the drain (check local regulations) |
| Empty pouch | Solid waste |
| Cassette | Solid waste or recycle (if accepted) |
| If you have... | Do this |
|---|---|
| Unopened pouches | Return to 4°C refrigerator |
| Opened pouch (1 gel used) | Reseal if possible; use remaining gel within 1 week |
| Opened pouch (all gels used) | Discard pouch |
| Step | Instructions |
|---|---|
| 1 | Rinse tank with distilled water |
| 2 | Wipe dry with paper towels |
| 3 | Remove any buffer residue (can cause corrosion) |
| 4 | Store tank upside down to dry |
Print this page and keep it at your bench.
Before starting:
Warm gel to RT (5–10 min, pouch closed)
Prepare fresh 1X running buffer
Boil samples (95°C, 5–10 min)
Step 1: Prepare gel
Open pouch, remove cassette
Remove comb (slowly, straight up)
Remove bottom tape
Rinse wells (syringe + buffer, 2–3× per well)
Step 2: Set up tank
Insert gel (short plate inward)
Fill inner chamber (cover wells)
Fill outer chamber (to line)
Check for leaks
Step 3: Load samples
Use gel-loading tips
Load ladder (5–10 µL, lane 1)
Load samples (10–30 µL)
Balance empty lanes with loading buffer
Step 4: Run gel
120V constant
Run until dye front reaches bottom (45–60 min)
Step 5: Remove gel
Open cassette with spatula
Cut off stacking gel
Proceed to transfer or staining
Running conditions quick reference:
| Voltage | Time |
|---|---|
| 120V constant | 45–60 min |
| 80V (stacking) → 150V (resolving) | 20 min + 30–40 min |
A: Yes, absolutely. This is the most skipped step and the #1 cause of problems. Rinsing removes unpolymerized acrylamide and air bubbles. Just do it.
A: 5–10 minutes at 95°C. 5 minutes is minimum. 10 minutes is better for difficult samples (membrane proteins, aggregates).
A: 120V constant works well for most applications. For sharper bands, use 80–100V for stacking (first 20 min) then 150V for resolving.
A: Stop when the dye front (blue line) reaches the bottom of the gel. If your smallest protein is >25 kDa, you can stop a few minutes earlier.
A: Not recommended. Reused buffer causes smearing and slower runs. Use fresh buffer for each run.
A: Possible causes:
Old running buffer → make fresh
Buffer concentration wrong (should be 1X)
Low voltage → check power supply settings
Inner chamber leak → check and reseat gel
A: Possible causes:
Buffer concentration wrong (too dilute)
Voltage too high → reduce to 120V
Gel percentage too low for your proteins
A: Not necessary for Ahelixbio pre-cast gels. Rinsing the wells removes unpolymerized acrylamide, so pre-running is optional.
A: Yes. Place one gel on each side of the inner chamber. Run conditions are the same.
A: Reseal the pouch with tape and store at 4°C. Use within 1 week. Better yet, only open pouches when you're ready to use all gels inside.
A: Ladder is purified protein and runs cleanly even with poor technique. Sample smearing usually means:
Too much protein loaded
Samples not fully denatured
Protease activity in lysate
A: Yes. Use Coomassie Blue, Silver stain, or any standard protein stain. Ahelixbio gels stain like any other SDS-PAGE gel.
A: Try these:
Run at lower voltage (80–100V)
Use fresh running buffer
Reduce protein load (20 µg instead of 50 µg)
Boil samples longer (10 min)
Chill buffer with ice bath
A:
1.0 mm: Sharper bands, uses less antibody, lower sample volume
1.5 mm: Can load more sample, better for low-abundance proteins
A: Yes. Ahelixbio gels have low autofluorescence and work with LI-COR and other fluorescent Western blot systems.
| Step | Critical Point |
|---|---|
| 1. Warm gel | 5–10 min at RT (pouch closed) |
| 2. Remove comb | Slowly, straight up |
| 3. Rinse wells | 2–3× per well with buffer (DON'T SKIP) |
| 4. Load samples | Use gel-loading tips, expel slowly |
| 5. Run at 120V | Until dye front reaches bottom |
| 6. Remove gel | Cut off stacking gel |
| 7. Transfer | Check with Ponceau S |
Using Ahelixbio pre-cast gels is straightforward once you know the steps. The two most important things to remember:
Rinse the wells – This prevents 90% of problems
Use fresh running buffer – Old buffer causes smearing
Follow this protocol, and you'll get sharp, reproducible bands every time.
Print the quick protocol card and keep it at your bench. You'll thank yourself later.
Have questions about using Ahelixbio pre-cast gels? Drop them in the comments below.
