Molecular Lab Teaching Program Outline
This teaching program introduces learners to essential molecular laboratory concepts, safe laboratory practice, and core experimental techniques used in modern biological, clinical, and biotechnology settings.
1. Program Goals
- Build foundational understanding of DNA, RNA, genes, and molecular workflows.
- Develop safe and accurate laboratory habits, including biosafety, contamination control, and documentation.
- Train learners in key techniques such as pipetting, nucleic acid extraction, PCR, gel electrophoresis, and data interpretation.
- Strengthen scientific reasoning through experimental design, troubleshooting, and reporting.
2. Target Audience and Prerequisites
The program is designed for undergraduate, clinical laboratory, or biotechnology trainees. Recommended prerequisites include introductory biology, basic chemistry, and familiarity with scientific measurement and laboratory safety principles.
3. Program Structure
- Orientation and Safety: laboratory rules, PPE, biosafety levels, waste handling, and contamination prevention.
- Core Molecular Biology Concepts: DNA/RNA structure, base pairing, gene expression, enzymes, and molecular diagnostics workflow.
- Laboratory Skills Bootcamp: micropipetting, solution preparation, serial dilution, labeling, and lab notebook practice.
- Nucleic Acid Extraction: sample preparation, extraction methods, quality checks, and storage considerations.
- PCR and Amplification: primer basics, reaction setup, thermal cycling, controls, and endpoint PCR analysis.
- Gel Electrophoresis: gel preparation, sample loading, DNA ladder use, visualization, and interpretation.
- Quantitative and Applied Methods: introduction to qPCR, RT-qPCR, sequencing concepts, or molecular diagnostics case studies.
- Capstone Investigation: student-designed or instructor-guided project integrating extraction, amplification, analysis, and reporting.
4. Teaching Schedule
| Week | Focus | Primary Activity |
| 1 | Orientation and biosafety | Safety briefing, PPE practice, contamination-control simulation |
| 2 | Foundational molecular biology | DNA/RNA concept workshop and workflow mapping |
| 3 | Essential lab skills | Pipetting accuracy, dilution practice, lab notebook setup |
| 4 | Nucleic acid extraction | DNA extraction and quality assessment |
| 5 | PCR | PCR setup, controls, and thermal cycling overview |
| 6 | Gel electrophoresis | Gel running, visualization, and band interpretation |
| 7 | Applied molecular methods | Case study in diagnostics, qPCR, sequencing, or gene expression |
| 8 | Capstone and communication | Final project presentations and lab report submission |
5. Assessment Strategy
- Pre-program diagnostic quiz to assess baseline understanding.
- Skills checklists for pipetting, safety practice, and sample handling.
- Laboratory notebook reviews for accuracy, completeness, and reflection.
- Data interpretation exercises using PCR and gel electrophoresis results.
- Final capstone report or presentation demonstrating experimental reasoning.
6. Resources and Materials
- Personal protective equipment, laboratory notebooks, and standard operating procedures.
- Micropipettes, tips, tubes, racks, centrifuge access, and basic molecular reagents.
- DNA extraction kits or classroom-safe extraction materials.
- PCR reagents, primers, thermal cycler access, agarose gel materials, and visualization equipment.
- Sample datasets for learners who cannot access all wet-lab equipment.
7. Implementation Notes
Instructors should adapt the program to local biosafety requirements, available equipment, learner level, and institutional policies. Where wet-lab access is limited, virtual simulations, instructor demonstrations, or prepared datasets can preserve the learning objectives while reducing logistical barriers.
8. New members Onboarding Guide
This section is adressed to new members joining the Inmunology and Molechular Biology Laboratory, and covers the practical, procedural and cultural foundations every incoming member must masrte before working independently on the bench.
8.1 Lab Policies and Hierarchy
The lab follows a mentorship-based hierarchy inspired by the traditional kohai-sempai-sensei model. This structure exists to guarantee that every new member has a clear point of contact for training, supervision, and troubleshooting, and to preserve consistent technique and safety standards across generations of lab members.
- Kohai (Junior): newly joined trainees. Kohai requires supervision to realize any work, as run reagents from share stock, or operate shared equipment (thermal cyclers, qPCR machines, centrifuges) until formally signed off by a sempai.
- Sempai (Senior): experienced members responsible for direct, hands-on training of one or more kohai. Sempai co-sign kohai notebooks, approve competency checklists, and are the first point of contact for technical questions.
- Sensei (Professor): Lab’s scientific direction, aprrove protocols and budget for reagents/bookings, resolve escalated issues, and have final autority on safety and data-integrity matters
General policies:
- Evey kohai is assigned as a sempai mentor during onboarding and must complete a documented competency checklist (pipetting, extraction, PCR, qPCR) before working independently.
- Attendance, use of PPE (gloves, lab, coat, closed-toe shoes, and eye or respiratory protection in necessary cases) and adherence to biosafety level requirements are always mandatory.
- No food, drink, or personal electronics are permitted at the bench. Long hair must be tied back and jewelry that could contaminate samples must be removed.
- All members must sign in/out of shared equipment logbooks and report any incident, spill, or equipment malfunction immediately to the sempai on duty or the sensei.
- Respectful, collaborative communication across hierarchy levels is expected; the kohai-sempai-sensei structure is a support system, not a rank of authority for personal matters.
8.2 Laboratory Work Protocol
At Lab’s manners and respect are fundamental. Before starting any bench work, new members must follow the Lab’s standard operating sequence:
- Review the day’s experimental plan with the assigned sempai and confirm reagents, equiptmen, and booking slots are available.
- Don PPE, Disinfect the bench surface with 70% ethanol (or designated disinfectant), and organize a clean workspace before opening any reagents or samples.
- Label all tubes, plates, and solutions clearly with sample ID, date, and initials before use, following the Lab’s labeling convention.
- Record every step, reagent lot number, volume, and observation in the laboratory notebook in real time; not from memory afterward.
- Segregate and dispose of biological chemical, and sharps waste according to the posted waste-stream protocol; never mix waste types.
- Decontaminate the bench, return shared equipment to its designated location, and log equipment to use at the end of every session.
- Report deviations from protocol, contamination events, or unexpected results to the sempai/sensei before repeating or continuing the procedure.
8.3 Use of micropipettes
Accurate pippetting is the single most important hands-on skill for a new member and is assessed before any independent bench work is authorized. Furthermire you can review the Lab’s video to complement your knwledge about it
- Select the pippette whose range matches the targent volume; never set a volume outside the pipette’s rated range (e.g. do no set a P20 above 20 µL).
- Use the correct tip for each pipette model and always attach a fresh tip for each new reagent or sample to prevent cross-contamination.
- Use the forward pipetting technique for standard aliquoting (press to the firts stop to aspirate, firts stop to dispense) and the teverse pipetting technique for viscous or foaming liquids (e.g., glycerol-containing buffers, enzymes).
- Hold the pipette vertically when aspirating, inmerse the tip only 2-4 mm below the liquid surface, and pipette slowly and consistently to avoid air bubbles and volume error.
- Never lay a loaded pipette on its side, never let go of a pressed plunger while the tip is submerged, and never use a pipette without a tip attached.
8.4 Volumes, Calculations, Solutions, Booking, and Reagent Handling
New members must be comfortable performing routine laboratory calculations before handling reagents independently:
- Concentration and dilution math, including the C1V1 = C2V2 relationship for preparing working solutions from stocks, and serial dilution design.
- Molarity, percentage (w/v and v/v), and unit-conversion calculations (ng/µL, µM, mM) commonly required for nucleic acid and buffer preparation.
- Master mix scaling: calculating per-reaction and total reaction volumes, including overage (typically 5–10%) to compensate for pipetting loss.
Solutions and reagents:
- All buffers and stock solutions must be prepared following the written Kardex or SOP, labeled with concentration, preparation date, preparer’s initials, and expiration/storage conditions.
- Reagents are stored at their required temperature (room temperature, 4°C, –20°C, or –80°C) and returned immediately after use; freeze-thaw cycles must be minimized and logged for sensitive reagents (enzymes, master mixes, RNA).
- Any reagent running low, expired, or behaving unexpectedly must be reported to the sempai/sensei so it can be reordered or requalified.
Equipment and space booking:
- Shared equipment (thermal cyclers, qPCR machines, centrifuges, biosafety cabinets, spectrophotometers) must be reserved through the Lab’s booking system or shared calendar.
- Bookings should include estimated start/end time and be canceled promptly if no longer needed, to keep equipment available for other members.
- Priority conflicts are resolved by the senpai on duty or escalated to the sensei; time-sensitive experiments (e.g., live samples, thawed reagents) take precedence.
8.5 Nucleid Acid Extraction Protocol
Tihs procedure covers DNA purification with PCI, DNA/RNA precipitation, RNA extraction with TRIzol, and DNA recovery from agarose gel by column purification.
Biosafety Conditions
Biosafety is preventive and mandatory before starting any extraction, to avoid short-term accidents (chemical burns, environmental contamination) and long-term risks (neuropathy, carcinogenicity) associated with phenol and chloroform.
- Wear high-quality gloves (preferably new), a long-sleeve lab coat, and closed laboratory shoes; fully cover any wounds to minimize contact with susceptible tissue.
- Perform every step involving phenol or chloroform inside a Class II biosafety cabinet, following the cabinet’s operating instructions.
- Stop the procedure immediately if any adverse reaction or discomfort occurs, and notify the Lab director or person in charge if necessary.
- Minimum PPE for this protocol: gloves, white long-sleeve lab coat (fluid-resistant preferred), and closed laboratory shoes.
Equipment
- Refrigerated microcentrifuge.
- Adjustable or fixed-volume micropipettes, matched to the volumes required by the protocol.
Materials and Reagents
Materials:
- Sample, latex or nitrile gloves, micropipette tips, timer/stopwatch, 1.5 mL or 2 mL vials, GenElute Agarose Spin Columns kit.
Reagents:
- PCI (phenol-chloroform-isoamyl alcohol), TRIzol, absolute ethanol (99%), 75% ethanol, TE buffer or autoclaved deionized distilled water.
A. DNA Purification with PCI (Phenol-Chloroform-Isoamyl Alcohol) + Ethanol Precipitation
PCI purification:
1. Place the sample to be purified in a 1.5 mL vial, in a volume no greater than 750 µL.
2. Add PCI (the phase at the bottom of the liquid) in a 1:1 ratio with the sample. Mix by inversion 5–7 times.
3. Centrifuge at 13,500 rpm, 4°C, for 10 min.
4. Collect the upper phase (supernatant) into a new vial, avoiding the lower phase to reduce sample contamination.
DNA precipitation:
1. Add 3 M sodium acetate, 1/10 of the sample volume. Mix by inversion 3–4 times.
2. Add cold (–20°C) 99% ethanol, 2 to 2.5 times the sample volume. Mix by inversion 3–4 times.
3. Centrifuge at 15,000 rpm for 15 min at 4°C. A pellet containing the precipitated DNA should be visible at the bottom of the vial.
4. Discard the supernatant by inverting the vial over absorbent paper or a paper towel. Do this only once and let the tube drain for a maximum of 5 min; once fully drained, return the vial to its upright position (do not invert again, or the pellet may detach).
5. Add 1 mL of cold (–20°C) 75% ethanol. DO NOT VORTEX OR MIX.
6. Centrifuge at 15,000 rpm for 10 min at 4°C.
7. Discard the supernatant by inversion, as described above.
8. Remove any residual supernatant with a 10 µL pipette, taking care not to disturb the pellet.
9. Resuspend and dissolve the pellet in autoclaved deionized distilled water or TE buffer (check the PCR protocol or the Lab’s master solutions list).
B. RNA Extraction with TRIzol
1. Aliquot 1000 µL of TRIzol into a 2 mL vial.
2. Place the tissue or sample in the tube (depending on sample size, it may be distributed across several vials).
3. Homogenize the tissue using a homogenizer.
4. Add 200 µL of chloroform (TRIzol:chloroform ratio = 5:1). Mix by vortex or inversion. Incubate at 4°C for 5 min.
5. Centrifuge at 15,000 rpm for 15 min at 4°C.
6. Transfer the supernatant with a pipette to a 1.5 mL vial. Add isopropanol (1:1 with the volume in the vial). Mix by inversion and incubate at –20°C for 15–30 min (samples may be kept at –80°C overnight).
7. Centrifuge at 15,000 rpm for 15 min at 4°C (a yellow or white pellet should be visible at the bottom of the vial). Discard the supernatant by inversion ONCE, onto a paper towel, without turning the vial back and forth.
8. Add 1 mL of cold (–20°C) 75% ethanol.
9. Centrifuge at 15,000 rpm for 15 min at 4°C. Discard the supernatant by inversion ONCE and remove all remaining ethanol using a 10 µL micropipette.
10. Resuspend the pellet in DEPC-treated water (ultrapure water with diethyl pyrocarbonate). Add the water and gently tap the bottom of the vial by hand until the pellet dissolves.
11. Measure RNA concentration using a NanoDrop or another method (e.g., Qubit).
12. Store at –20°C until use.
C. DNA Extraction from Agarose Gel by Spin-Column Purification
1. After running gel electrophoresis and identifying the band(s) to purify, place the gel on plastic wrap (“vinipel”).
2. Cut out the band(s) of interest with a scalpel under UV light, in a fully darkened room.
3. Prepare 1 purification column and 2 tubes (from the GenElute Agarose Spin Columns kit).
4. Place the purification column into one of the tubes.
5. Add 100 µL of TE buffer.
6. Centrifuge at 15,000 rpm for 10 seconds at room temperature.
7. Change the tube and place the purification column into a new tube.
8. Add the gel product (the previously excised band) to the purification column. Cut the gel into smaller pieces or pass it through an insulin syringe to help it pass through the column and increase DNA recovery.
9. Centrifuge at 15,000 rpm for 15 min at 4°C.
10. Discard the column and calculate the volume of the filtrate. If the volume is less than 100 µL, add sufficient autoclaved deionized distilled water to reach that volume.
11. Add 3 M sodium acetate, 1/10 of the sample volume. Mix by inversion 3–4 times.
12. Add cold (–20°C) 99% ethanol, 2 to 2.5 times the sample volume. Mix by inversion 3–4 times.
13. Centrifuge at 15,000 rpm for 15 min at 4°C. A pellet containing the precipitated DNA should be visible at the bottom of the vial.
14. Discard the supernatant by inverting the vial over absorbent paper. Do this only once and let the tube drain for a maximum of 5 min; return the vial upright once fully drained (do not invert again).
15. Add 500 µL of cold (–20°C) 75% ethanol. DO NOT VORTEX OR MIX.
16. Centrifuge at 15,000 rpm for 10 min at 4°C.
17. Discard the supernatant by inversion, as described above.
18. Let it air-dry for a maximum of 5 min and remove any residual supernatant with a 10 µL pipette, taking care not to disturb the pellet.
19. Resuspend and dissolve the pellet in autoclaved deionized distilled water or TE buffer (check the PCR protocol or the Lab’s master solutions list).
20. Store at –20°C until use.
8.6 PCR (Polymerase Chain Reaction)
This procedure reproduces the Lab’s official SOP, “Protocolo de Reacción en Cadena de la Polimerasa (PCR): Punto Final, PCR en Tiempo Real (qPCR) y Electroforesis Horizontal en Gel de Agarosa” (Laboratorio de Inmunología y Biología Molecular, LIBM), adapted here for new-member training on conventional (endpoint) PCR.
Biosafety Conditions
- Wear gloves (latex, nitrile, or vinyl — preferably new), a long-sleeve lab coat, and closed laboratory shoes; fully cover any wounds to minimize contact with susceptible tissue.
- Stop the procedure immediately if any adverse reaction or discomfort occurs and notify the Lab director or person in charge if necessary.
- Minimum PPE for this protocol: gloves, white long-sleeve lab coat (fluid-resistant preferred), and closed laboratory shoes.
Equipment
- Thermocycler — ProFlex™ PCR System (Applied Biosystems, Carlsbad, CA, USA).
- Micropipettes, matched to the volumes required by the protocol.
- Vortex.
- 1.5 mL microcentrifuge tubes.
- 200 µL microcentrifuge (PCR) tubes.
Materials and Reagents
Materials:
- Cold station (e.g., CoolCaddy), latex or nitrile gloves, micropipette tips, micropipettes of different volumes, 1.5 mL and 200 µL vials (for PCR), 100 µL tubes (for qPCR), permanent marker.
Reagents:
- DNA sample (genomic or complementary), deionized distilled (molecular-grade, Type 1) water, Taq polymerase buffer, dNTPs, primers specific to the target (forward and reverse), magnesium chloride (MgCl₂), Taq polymerase enzyme.
Note: some commercial suppliers provide pre-made master mixes; in that case, follow the manufacturer’s recommendations or the Lab’s in-house standardization instead of the component list below.
Conventional (Endpoint) PCR Procedure
1. Label the tubes with each sample’s identification before starting the experiment.
2. Calculate the required volume of each reagent for 1 reaction, bringing the final volume to 25 µL with deionized distilled water (dd water).
3. Multiply each volume by the number of samples to be run to prepare the Master Mix (it is recommended to calculate for one extra reaction to compensate for pipetting loss).
4. Table 1 shows an example Master Mix calculation for 4 samples, based on the suggested protocol for GoTaq® G2 Flexi DNA Polymerase (Promega, USA):
| Component | Volume (1 reaction) | n | Total |
| dd water | 14.875 µL | 5 | 74.375 µL |
| Buffer 5x | 5 µL | 5 | 25 µL |
| dNTPs (1.5 mM) | 1 µL | 5 | 5 µL |
| Primer F (10 µM) | 1 µL | 5 | 5 µL |
| Primer R (10 µM) | 1 µL | 5 | 5 µL |
| MgCl₂ (25 mM) | 1 µL | 5 | 5 µL |
| Taq polymerase | 0.125 µL | 5 | 0.625 µL |
| Template | 1 µL | – | – |
| Total | 25 µL | – | 120 µL |
Table 1. Components and volumes of a Master Mix (25 µL) for 4 samples.
5. Prepare the Master Mix by adding the reagents in the order shown in Table 1, starting with dd water and finishing with Taq polymerase (reagent order follows reagent cost/stability). Remove the Taq polymerase from the freezer only when ready to add it to the Master Mix and return it to –20°C immediately afterward.
6. Vortex and then centrifuge the master mix for 5 seconds, avoiding bubble formation.
7. Using a 10–100 µL micropipette, dispense 24 µL of Master Mix into each of the previously labeled 200 µL PCR tubes.
8. Add 1 µL of template (DNA sample) to each PCR tube containing the Master Mix.
9. Perform a quick spin-down centrifugation for 5 seconds, avoiding bubble formation.
10. Turn on the thermocycler and program the thermal profile appropriate to the sample. For example:
| Step | Temp (°C) | Cycles | Time |
| Pre-denaturation | 95 | 1 | 3 min |
| Denaturation | 95 | 35 | 30 sec |
| Annealing | 55 (variable) | 35 | 30 sec |
| Extension | 72 | 35 | 30 sec (variable) |
| Final extension | 72 | 1 | 5 min |
Table 2. Conventional PCR thermocycler run conditions.
a. Place the PCR tubes in the thermocycler and start the run.
Notes
- PCR component amounts vary depending on the reagent brand used (e.g., some Taq formulations already contain MgCl₂ and do not require it to be added separately to the master mix).
- Thermocycler conditions vary by sample: for example, some Taq enzymes polymerize approximately 500 bp every 30 seconds, so a 300 bp fragment needs about 30 seconds of extension while a 900 bp fragment needs about 1 minute.
Primer annealing temperature depends on the primer melting temperature (Tm) and the sample being analyzed and must be verified individually for each experiment.