Outreach Science Ontario

Cloning genes is THE most fundamental technique in modern biotechnology. Before a gene can be used as a tool to produce large quantities of a protein (that might be needed for use as a drug or vaccine) or before it can be used to genetically modify an organism (GMO), it must first be cloned.
This module is designed to demonstrate methods for cloning your favourite gene, starting with a limited sample. We use polymerase chain reaction (PCR), a protocol developed in the mid 1980s, to amplify your favourite gene in a cloning process. After PCR, we verify that the correct gene has been amplified using a portion of the reaction products on an agarose gel using electrophoresis. The amplified DNA is also ligated directly into a standard cloning vector and transformed into bacteria that replicate the recombinant plasmids to a high copy number. Transformed bacteria colonies will be isolated and the presence of the insert DNA verified by restriction mapping.

We have chosen to clone a gene (KanR) encoding a protein which allows bacteria to grow in the presence of the antibiotic kanamycin. We have supplied bacterial DNA from a strain expressing this gene to serve as the template for the PCR.

N.B. Full protocols are given in the Student Instructions (this is a Word file). Daily setup and safety instructions for teachers are provided below.

General:
I - Kit Storage:
1. Store reagents (PCR and T-vector ligation kits) at –20C.
2. Store competent cells (large Styrofoam box) at -20C and leave sealed until ready to use.
3. Just before use, thaw the materials at room temperature, then place them immediately on ice.
4. Plates are best stored in the fridge, but can be left at room temperature.
5. All other equipment and reagents are room temperature.

II - General Instructions:
1. There are enough reagents for 10 workgroups (with a bit of spare) to complete the following protocol.
2. Ensure that all students read the directions carefully and are particularly aware of flame and sharps hazards.
3. Ensure that students use a new pipette tip each time they go into a reagent. Have students practice pipeting techniques using water dropped onto paper toweling to check for accuracy. Measure the circles, ensuring each is the same on all practice runs. Have them compare their results with other students.
4. Pipettes, tips, microtubes, and waste beakers are needed every day; see below for additional equipment needed for each procedure.
5. Place any waste in plastic containers and return it with the kit in the biohazard bag or a garbage bag.

Have several small buckets of ice (preferably crushed) for the PCR reagents and get out two PCR reagents (template + primer mix, PCR reaction mix) just prior to class
Specific Equipment: PCR machine, smaller microtubes
Multi-coloured permanent markers are useful for labeling the small microtubes

An assembly-line setup works well, as there is usually only one tube of each PCR reagent provided.
Make sure assembled reactions are kept on ice until all are ready to be placed in PCR machine.
Check the machine a few minutes after starting the program to ensure it is temperature cycling.
The agarose gel for Day 2 can be prepared today and stored in the fridge wrapped in saran wrap if desired.

The UV transilluminator will cause intense skin burns and eye damage if turned on without the shield in place. Ensure that the shield is properly in place before use. Note also that the glass plate is easily broken. Handle with care.

The gel takes ~30 minutes to solidify, so prepare prior to class.
Specific Equipment: gel apparatus and transilluminator.
Specific Reagents: Agarose, Sybr safe DNA gel stain, 10X TBE (teacher use only), loading dye, DNA marker. All are room temperature reagents, except the DNA marker (freezer).

Make sure students add the loading dye to their samples prior to loading and that they record which well their sample is in.
Make sure the DNA marker is loaded (and has loading dye added as well).
The DNA marker is for determining the size of your sample DNA fragments and will have many bands. Student samples should show one band.
The loading dye is not toxic but can stain hands and clothing (usually washable).
The image (right) shows the DNA marker as it will appear on your gel with sizes of the DNA fragments labelled.

Have several small buckets of ice (preferably crushed) for ligation reagents.
Ligation buffer and vector can be thawed, mixed by flicking, and placed on ice prior to class, but leave ligase enzyme in freezer until ready to use.

The assembly-line again works well. Make sure the ligase is the last stop (last reagent added).
Be sure to emphasize the small volumes and setting the pipettes properly (only 1 ul of ligase), as students may use too much and there is only enough for 10-12 reactions.
If a student does use too much (e.g., adds 10 ul instead of 1 ul of a reagent), the reaction can be scaled up with the other reagents, e.g., make a 10X mix like you would make 10X of a recipe. This way, there will be enough for everyone to use for the transformation step.
If there doesn’t appear to be enough of a reagent, spin the tube (with a balancer tube) in the centrifuge for 10 seconds.

Cell spreaders: BE SURE students seal the end of the glass pipettes COMPLETELY so ethanol will not get in when the spreader is dipped as this is very dangerous when the ethanol is burned off!!
Make sure students do not go near the jar of ethanol with a flaming spreader as they could light it on fire. If this happens, quickly put the lid on to put out the flame.

Have several beakers of crushed ice for the competent cells. Leave cells at -20C until ready to use.
Also note that the EcoRI enzyme is stored with the competent cells and should be removed once the box is opened and kept at -20C.
Prepare a 42C water bath and a 37C incubator.
Specific equipment needed: Centrifuge (set at 4000 rpm), tube floater, cell spreaders (glass pipettes), bunsen burner.
Specific reagents needed: student’s ligation reactions, competent cells, SOC, MacConkey plates (red), ethanol jar.
MacConkey plates should be warmed to room temperature before use.
You may wish to make a cell spreader prior to class for practice and example.

There are several sets of the transformation reagents, so stations, each with one set of reagents and 1-2 groups of students, work well.
Longer incubations at 37C (after adding SOC) will improve the transformation, so leave as long as possible.
Wait until all students have finished the 10 minute incubation on ice (longer is fine); place all tubes in the floater on ice and heat shock them all at once.
Place the ethanol jar on the opposite side of the Bunsen burner from the plates. This way, there is less chance of students holding the flaming spreader near the jar.

Day 4: Miniprep and Restriction Digest
Safety: No special concerns.
Setup:

Just prior to class, add 1 ml of sterile water to the lysozyme (in a 1.5 ml tube in the freezer) and vortex to dissolve the powder. Keep on ice (only reagent that needs ice today).
Prepare a boiling water bath.
Specific Equipment needed: Vortex, toothpicks, centrifuge (return setting to 13000 rpm), and floater, 5 ml plastic pipettes, and green pipettes for the isopropanol.
Specific Reagents: STET buffer, isopropanol, 70% ethanol, and sterile water (all found in the 5 ml tube rack). Lysozyme, RNase, and cell pellets (freezer). For the digest: 10X enzyme buffer, EcoRI (freezer).

Except for the lysozyme, there are several sets of the reagents, so a station setup works well.
The lysozyme is a fairly stable enzyme and can be kept on ice during class.
EcoRI should be left in the freezer until ready to use and then kept on ice during use.
Remind students to OPEN the lids of their tubes before boiling and to balance samples in the centrifuge.
DNA pellets can be very small and difficult to see, so students need to be careful during ethanol removal to avoid removing the pellet.

Day 5: Analyzing Digests by Gel Electrophoresis and Phenotype analysis
SAFETY and Setup:

See Day 2 above
On this gel, students should see two bands: the larger one is the original vector (closest to the gel wells), the smaller one is the KanR insert.
If they see only one band it indicates their digest was unsuccessful.
A large diffuse band (RNA) indicates the student didn’t add RNase.