Laboratory Fundamentals Protocol

How to access MSDS:

1. Check the reagent bottle label for manufacturer information
2. Search online: "[Chemical name] MSDS [Manufacturer]"
3. Consult your institution's chemical inventory system
4. Request from laboratory instructor or chemical supplier

PPE Best Practices:

• Inspect gloves for tears before use
• Change gloves between different procedures or when contaminated
• Remove lab coat before leaving laboratory
• Never wear PPE in public areas (cafeteria, library, etc.)
• Dispose of contaminated PPE properly

Critical practices:

• Change gloves between different procedures or when contaminated
• Never touch face, hair, phone, or notebook with gloved hands
• Keep tube caps closed except when actively pipetting
• Never set pipette tips on bench surface
• Use filter tips for DNA work
• Work in one consistent direction (prevents cross-contamination)
• Label all tubes before adding samples

The C₁V₁ = C₂V₂ Formula

Where:

• C₁ = initial (stock) concentration
• V₁ = volume of stock needed
• C₂ = final (working) concentration
• V₂ = final total volume needed

Rearranging to solve for V₁:

V₁ = (C₂ × V₂) / C₁

Exercise 1: Buffer Dilution

You need to prepare 1X TAE running buffer from 50X stock for gel electrophoresis. You need 200 mL total.

Using C₁V₁ = C₂V₂:

• C₁ = 50X
• V₁ = ?
• C₂ = 1X
• V₂ = 200 mL

V₁ = (C₂ × V₂) / C₁ = (1 × 200) / 50 = 4 mL

Answer: Mix 4 mL of 50X TAE + 196 mL water = 200 mL of 1X TAE

Exercise 2: Primer Working Solution

You have 100 µM primer stock. Your protocol requires 10 µM working solution. Prepare 500 µL.

C₁V₁ = C₂V₂

• 100 µM × V₁ = 10 µM × 500 µL
• V₁ = 50 µL

Answer: Mix 50 µL stock + 450 µL nuclease-free water = 500 µL at 10 µM

Exercise 3: Serial Dilution

Create a 1:10 dilution series starting from 1 mg/mL stock. Prepare four dilutions, each 100 µL total.

• Tube 1: 10 µL stock + 90 µL water = 0.1 mg/mL (1:10)
• Tube 2: 10 µL from Tube 1 + 90 µL water = 0.01 mg/mL (1:100)
• Tube 3: 10 µL from Tube 2 + 90 µL water = 0.001 mg/mL (1:1000)
• Tube 4: 10 µL from Tube 3 + 90 µL water = 0.0001 mg/mL (1:10,000)

Aliquot Size Considerations:

• Calculate usage per experiment
• Plan for 3-5 uses per aliquot
• Account for pipetting dead volume (~5 µL for 1.5 mL tubes)
• Example: If using 2 µL per reaction, 4 reactions per week, aliquot 50 µL (allows for 20+ reactions, accounting for dead volume)

Aliquoting Procedure:

1. Label all aliquot tubes before starting (include name, reagent, concentration, date)
2. Keep stock and aliquots on ice during procedure
3. Mix stock thoroughly before aliquoting (invert 10-15 times or vortex gently)
4. Dispense into labeled tubes
5. Seal all tubes
6. Return stock to storage immediately
7. Store aliquots at appropriate temperature

Step-by-step procedure:

1. Select appropriate pipette for your volume range
2. Adjust volume using the adjustment knob
3. Attach tip by pressing pipette into tip firmly (should seal completely)
4. Pre-wet the tip (optional but recommended for accuracy):
   • Draw up and expel liquid 2-3 times
   • Ensures consistent liquid film inside tip
5. First stop technique (standard pipetting):
   • Hold pipette vertically
   • Press plunger to first stop (resistance point)
   • Insert tip 2-3 mm into liquid
   • Slowly release plunger to draw liquid
   • Wait 1 second for liquid to enter tip completely
   • Remove tip from liquid (still against tube wall)
   • Touch tip to destination tube wall
   • Press to first stop to dispense
   • Press to second stop (blowout) to expel remaining liquid
   • Remove pipette while keeping plunger depressed
6. Eject tip into waste container using tip ejector button

Procedure:

1. Press plunger to second stop (not just first)
2. Draw liquid (will over-aspirate)
3. Dispense by pressing to first stop only
4. Retain excess liquid in tip
5. Eject tip with excess liquid

Exercise 1: Serial Dilution with Visual Confirmation

Objective: Create a 1:2 serial dilution series to practice accurate pipetting and understand dilution factors.

Procedure:

1. Receive PCR strip tube with 100 µL colored "stock" in well 1
2. Add 50 µL water to wells 2-8
3. Transfer 50 µL from well 1 to well 2, mix by pipetting up and down 3-5 times
4. Transfer 50 µL from well 2 to well 3, mix
5. Continue through well 8
6. Observe color gradient (should decrease consistently)

Expected result: Each well should be approximately half the intensity of the previous well.

If gradient is inconsistent: Pipetting technique error. Practice with instructor supervision.

Exercise 2: Working Solution Preparation

Scenario: You have a "10X enzyme buffer" (dark blue dye). Prepare three working solutions:

1. 1X buffer (for PCR): 10 µL stock + 90 µL water = 100 µL
2. 0.5X buffer (for restriction digest): 5 µL stock + 95 µL water = 100 µL
3. 2X buffer (for master mix): 20 µL stock + 80 µL water = 100 µL

Visual check: Solutions should show different color intensities correlating with concentration.

Exercise 3: Aliquoting Practice

Objective: Prepare five 20 µL aliquots from a master solution.

Criteria for success:

• All five aliquots should be identical color intensity
• Volume accuracy: each should be 20 µL ± 1 µL
• No bubbles in solution
• Consistent technique for all five

Instructor checkpoint: Demonstrate your aliquoting technique for verification.

Exercise 4: Mixing Techniques

Practice different mixing methods:

1. Pipette mixing: Draw up and expel liquid 5-7 times
2. Vortex mixing: Brief 2-3 second pulse on vortex mixer
3. Flicking: Hold tube at top, flick bottom sharply 5-10 times
4. Inversion: Invert tube 10-15 times (for larger volumes)

Compare effectiveness with colored dyes and discuss when each method is appropriate.

Before starting:

• Date and title of experiment
• Objective or hypothesis
• Complete protocol with modifications
• Reagent lot numbers and concentrations
• Equipment used

During experiment:

• Observations (color changes, precipitation, timing issues)
• Deviations from protocol
• Problems encountered and solutions
• Environmental conditions if relevant (temperature, humidity)

After experiment:

• Raw data (all measurements, even if they seem wrong)
• Calculations
• Results summary
• Interpretation and conclusions
• Questions for follow-up

Example Entry Format:

Date: October 30, 2025
Experiment: DNA Extraction Comparison - Magnetic Bead Method
Objective: Compare DNA yield from mosquitoes preserved by three methods

Protocol: BioDynami HMW Genomic DNA Extraction (Automation Method)
Modifications: None

Samples:
- Sample 1: Frozen -80°C (received from instructor 2:05 PM)
- Sample 2: 95% ethanol (blotted dry before processing)
- Sample 3: Silica gel (appeared well-desiccated)

Reagents:
- TS Buffer, Lot #BD2024-10
- Proteinase K, Lot #BD2024-08 (stored -20°C)
- Lysis Buffer, Lot #BD2024-09

Observations:
- 2:15 PM: Tissue grinding complete, all samples homogenized well
- 3:00 PM: Lysis started, tubes in heat block at 56°C
- 3:45 PM: Sample 1 completely clear, Samples 2 and 3 slightly turbid

Results:
| Sample | Preservation | Qubit (ng/µL) | Total DNA (ng) |
|--------|-------------|--------------|----------------|
| 1      | Frozen      | 12.4         | 1240          |
| 2      | Ethanol     | 8.7          | 870           |
| 3      | Silica      | 5.2          | 520           |

Conclusions:
Frozen preservation yielded 2.4x more DNA than silica gel method.
Results consistent with class discussion predictions.

Next steps:
Use Sample 1 for PCR amplification next week.

Best Practices:

• Keep enzymes on ice or in ice block during use
• Pre-chill centrifuge for cold spins
• Allow frozen samples to thaw on ice (not at room temperature)
• Return temperature-sensitive reagents to storage immediately
• Verify heat block/thermal cycler temperatures before use
• Allow reactions to reach room temperature before opening (prevents condensation)

DNA/RNA work:

• Use filter tips for PCR and qPCR
• Separate pre-PCR and post-PCR work areas physically
• Never bring amplified DNA into reagent preparation area
• Change gloves frequently
• Use dedicated pipettes for different work areas if possible
• Clean pipettes regularly with DNA decontamination solution

Sample organization:

• Create clear physical separation between samples
• Process negative controls alongside experimental samples
• Include a "no template control" for PCR
• Label everything before starting
• Use tube racks to maintain sample order
• Never touch tube rims or caps with gloved hands

Technique:

• Brief "spin down": 2-3 seconds to collect liquid at tube bottom
• Microcentrifuge speeds: typically 12,000-16,000 × g
• For pelleting: ensure adequate time at maximum speed
• For separations: follow protocol specifications exactly

Procedure:

1. Calculate total volume needed (number of reactions + 10% overage)
2. Mix all common components in single tube
3. Vortex and spin down master mix
4. Aliquot master mix to individual tubes
5. Add sample-specific components (template DNA, primers if different)

Solutions:

• Use low-retention tubes and tips
• Keep tubes closed when not in use
• Work quickly
• Use carrier molecules (glycogen, BSA) if appropriate
• Spin down frequently to collect liquid
• Pre-wet tips when pipetting viscous solutions
• Visual confirmation: hold tube to light to see meniscus

Before starting:

• All reagents prepared correctly?
• All tubes labeled?
• Protocol reviewed completely?
• Positive and negative controls included?
• Equipment functioning properly?

During experiment:

• Volumes look correct?
• Colors/clarity as expected?
• Timing on schedule?
• Controls behaving appropriately?

After experiment:

• All data recorded?
• Results make sense?
• Controls worked?
• Any unexpected observations documented?

Biological Waste:

• Tissues and biological samples → Biohazard bag
• Contaminated tips and tubes → Biohazard bag
• Bacterial cultures → Autoclave before disposal

Chemical Waste:

• Organic solvents → Labeled chemical waste container
• Heavy metals → Separate labeled container
• Mixed waste → Follow institutional guidelines

Sharps:

• Needles, scalpels, broken glass → Sharps container
• Never overfill sharps containers (max 2/3 full)

Regular Trash:

• Uncontaminated packaging
• Paper towels used for cleaning
• Empty reagent tubes (if confirmed non-hazardous)