The general procedure
Laboratory One: Safety, laboratory equipment and introduction to pipetting
Description: Designed as a hands-on training session that introduces all students to the
equipment in a modern Molecular Biology/Genetics laboratory. Future labs build
upon this initial training, such that students from diverse educational backgrounds
start with a level playing field. Each student practices using three single channel,
and two multi-channel pipettors (one manual and one electronic). Each student also
receives instruction using basic laboratory equipment such as micro-centrifuges,
centrifuges, a balance and programming a PCR machine.
Learning Objective: Students are given a first iteration of hands-on use of common
molecular biology laboratory equipment. The procedures are designed to give the
instructor the maximum amount of time to identify those students who need
immediate, one on one instruction.
Cost per student: $0.25
Laboratory Two: Collection of samples, extraction of genomic DNA and polymerase
chain reaction setup
Description: Students are taken outside and encouraged to select plants from a region
of campus that has several different plants. In addition, each student is required to
draw the area from which they took their sample and a digital picture is taken of the
plant. Each student then extracts DNA from their plants using a single tube
extraction protocol. Dilutions of these DNA extractions are used in PCR with
different RAPD primers (RAPD primers are used for PCR setup due to their
likelihood of success on unknown genomic DNA).
Learning Objective: This is the first step in getting the student personally involved in
learning molecular biology. The student picks their plant, draws the area that they
have chosen and extracts DNA from their plant. This is also the first chance to
actively use the skills introduced in lab 1 and to improve documentation skills.
Cost per student: $2.25
Laboratory Three: Electrophoresis of collected PCR samples, Design assigned gene
primers and practice for lab practical.
Description: Students load their RAPD PCR samples from lab 2, then design primers
for assigned gene(s) using protocol in Figure 1. Students are currently assigned
multiple genes to increase their probability of success.
Learning Objective: This is usually the first time that each student uses online scientific
research databases and tools. This is also the first time that students may have to
make a decision on what organism to use if there is not a representative gene from
soybean. This is the first reinforcement of the concept that genes can be highly
conserved between organisms. Students are encouraged to select eukaryotes if
soybean sequences are unavailable, but in many cases must select a prokaryote as
the source organism for their gene sequence. This protocol personally involves the
student in the research to be performed in the lab.
Cost per student: $16.00
Laboratory Four: Lab Practical Exam
Description: Placement of the lab practical at this point in the course assures the
instructor that the student is ready for the next phase of the project, which requires
competency in laboratory techniques and forces the student to learn proper
techniques earlier, rather than later.
Timing the practical after the primer design lab also allows additional time for the
primers to be ordered and delivered
Laboratory Five: Primer dilution and initial PCR test of Soybean and Pea
Description: Primers ordered from lab three are processed by re-suspension in TE, then
dilution to a working concentration of 10 mM. An initial test of these primers is
conducted on soybean and Pea DNA. The students are given a PCR protocol and
are required to program the PCR machines de-novo.
Learning Objective: Resuspension of primers, pipetting accuracy
Cost per student: $2.00
Laboratory Six: Electrophoresis of initial PCR test: PCR setup based on electrophoresis
results
Description: Students load PCR product from lab five. After electrophoresis, the
amplification profile is analyzed and based on these data, each student now begins
the independent study portion of their instruction. There are four possible outcomes
from the previous PCR setup (Figure 1).
The course can end at this time, or can continue by screening all functional primer
pairs on a larger population of organisms.
Learning Objective: Electrophoresis, post-translational modifications (product is larger
than predicted from EST sequence), gel loading and pipetting
Cost per student: $varies
Laboratory Seven: Electrophoresis of populations.
Description: Electrophoresis of PCR product from lab six. Collect and analyze data as
instructor desires (phylogeny trees, statistical analysis, etc)
Learning Objective:
Cost per student: $varies
Louisiana Tech
Name:_________________________ Date: ___________
BISC310 Fall 2008
Lab 1: Safety; Introduction to Pipetting
I understand that I am responsible for my own
personal safety and for the safety of others
in the lab. I will not deliberately perform actions
that may endanger myself or others.
__________________________
Signature
Pipettor practice:
Individually:
Fill your 96 well plate as shown;
Red food coloring
Green Food Coloring
250 ul
40 ul
Blue food coloring
6 ul
025
040
060
0250
040
0600
0250
40
60
Blue tips
yellow tips
Pipetman
Oxford
Finnpipette
yellow/white tips
Using the single channel pipettor, load 10 ul of blue dye into 12 wells of a pre-poured
agarose gel
Using the digital (non-electronic) 16 channel pipettor, load 10 ul of blue dye into 12 wells
of a pre-poured agarose gel
Electronic pipettors:
Press the grey oval button until the display looks like this:
DISP steps
Place the tips into the blue dye, then press the Grey button
(fluid is drawn into the tips)
6
10.0
ul
Place the tips into the wells in column 1, then press the
grey button ONCE; this dispenses 8ul into each well.
Continue to each column, dispensing 8 ul into each well. You will have to refill your tips
halfway through the plate.
Single Channel Pipettor Accuracy Verification
Pipettor #
Pipettor
Type
Test volume
uL
Test Weight
Error % (vol/weight)
______
P10 or 20
3
_________
___________
9
_________
___________
60
_________
___________
180
_________
___________
220
_________
___________
980
_________
___________
______
______
P200
P1000
Measuring Accuracy %:
If test weight is greater than test volume: (test weight – test volume) x 100
test volume
If test weight is less than test volume:
(test volume – test weight)
test volume
x 100
To conduct this test, place a small “weigh boat” on a fine scale. set your pipettor to the
appropriate setting above, then pipette (carefully) water into the weigh boat. It is easiest
to start with the smallest volume first, then “TARE” in between each measurement,
without emptying the weigh boat, progressively testing upwards in volume. A 2-3% error
is considered acceptable (and normal).
Louisiana Tech
Name:_________________________ Date: ___________
BISC310 Fall 2008
Lab 2: Random Sample DNA Extraction and PCR
Section _________
Partner names ____________________ ____________________
Collection Location ______________________
Summary: We are collecting random leaf samples from different plants on campus,
extracting DNA from these samples, then PCRing the DNA using RAPD primers. These
primers are short, 10 base pair fragments of DNA that allow the initiation of the PCR
amplification reaction wherever a complementary sequence occurs in your target DNA.
Because they are random, the probability is high that you will have successful PCR.
These PCR’d samples will be electrophoresed in lab next week.
Drawing of Collection Location
Picture of Source Plant
(Will be provided next lab)
Group
Sample Collection
___
1. Draw the location where leaf was collected, have a picture taken of your plant,
using the provided flashcard to write a large, legible label for the picture
___
2. Punch a leaf tissue disk into a Labeled 2.0 ml tube (Section#_Group #).
Collect only one leaf sample per group of three students.
___
3. Place the tube in the ice bucket
DNA Extraction
___
4. Add 100 ul Extraction solution (tube “E”) to the collection tube. Make sure
leaf disk is covered by extraction solution.
___
5. Incubate at 95o C for 10 minutes. CAUTION – TUBES WILL BE HOT!!
___
6. Add 100 ul dilution solution (tube “d”), then shake to mix, briefly centrifuge
Individual
Individual Dilution of DNA samples
___
7. Label a 1.5 ml tube with your section and name, adding a “d” (these will
contain your diluted DNA)
___
8. Add 45 ul of water (tube “W”) to each dilution tube
___
9. Add 5 ul DNA from the leaf sample tubes into the labeled dilution tube.
Individual PCR setup
___
___
___
___
10. Label 3 0.2 ml PCR tubes with your initials and #’s 1, 2, 3.
11. Add 3 ul water to each tube
12. Add 10 ul PCR mix (with green dye – tube “T”) to each tube
13. Add 2 ul of a different primer to each tube
Primers used: tube tube 1: _____
___
___
___
tube 2:______
tube 3:______
14. Spin down
15. Add 4 ul of diluted DNA (from steps 7-9) to each tube, (high and on the side)
16. Spin down, note on the PCR machine layout diagram where you placed your
samples; Program the PCR machine as follows:
Lid 105; Wait; Auto
95o
5 min
95 o
30 sec
40 o
60 sec
72 o
30 sec
Goto 2 Rep 40 times
72 o
5 minutes
Hold 4 o
Louisiana Tech
Name:_________________________ Date: ___________
BISC310 Fall 2008
Lab 4: Primer Dilution and Initial PCR Setup
Section ______________________
Summary: The primers that you ordered last week are shipped dried. You must first re-hydrate
them, then test them on a small subset of the pea and soybean parents. You will electrophorese
the PCR reactions from this week at the start of next weeks lab. If they are successful, you will
set up a much larger PCR screen of parental DNA with your primers.
YOU MUST WEAR GLOVES DURING THIS PROCEDURE
Re-hydration of Primers
___
1. Spin down the tubes to collect any lyophylized (dried) primer at the bottom of
the tube.
___
2, Add TE to each tube (10 X the nmol value)
Example: Add 301 ul TE to the primer shown
___
3. Make sure the cap is closed, spin down again and
place the primer tubes @ 60o for 60 minutes.
Dilution of Primers
___
4. Lightly shake your re-hydrated primers and spin them down
___
5. Label 4 1.5 ml tubes with the provided cryogenic labels (1 for each primer)
___
6. Add 450 ul Water to each dilution tube
___
7. Add 50 ul of your stock primer to the labeled dilution tube
Screen of four soybean and four pea lines
___
___
8. Add 30 ul WATER to the green master mix tube (this tube already contains
80 ul 2X TAQ master mix)
___
___
9. Add 10 ul of your DILUTED “L” primer to the tube with the PCR mix (with
green dye)
___
___
10. Add 10 ul of your DILUTED “R” primer to the tube with the PCR mix (with
green dye)
___
___
11. Spin down master mix tube
___
___
12. Add 15 ul of the green master mix from steps 8-11 into 8 PCR tubes provided
(tubes already contain soybean/pea DNA)
___
___
13. Spin down the tubes, label with your initials, close caps. Note on PCR
machine layout diagram where you placed your samples
Louisiana Tech
BISC310
Lab 5: PCR of Parents
Name:_________________________ Date: ___________
Section ______________________
YOU MUST WEAR GLOVES DURING THIS PROCEDURE
Soybean amplicons observed (16 reactions)
1
___
1. Label 16 0.2 ml PCR tubes (2 x 8 tubes w/caps) with your initials and sample number
(1-16)
___
2. Add 60 ul WATER to the green master mix tube labeled “1”
(this tube already contains 180 ul 2X Taq master mix)
___
3. Add 20 ul of your DILUTED “L” primer to the tube with the PCR mix (with
green dye)
___
4. Add 20 ul of your DILUTED “R” primer to the tube with the PCR mix (with
green dye)
___
5. Spin down master mix tube
___
6. Add 15 ul of the green master mix from steps 2-5 into the labeled tubes
from step 1.
___
7. Add 5 ul of diluted DNA into the 16 PCR tubes as shown below (we will add –
just let us know the tubes are ready)
tube 1
tube 2
tube 3
tube 4
tube 5
tube 6
tube 7
tube 8
84-1
4290
Harosoy
Clark
4895
Jackson
Anoka
A7
___
8. Spin down reactions
___
9. Place in PCR machine and note the location
tube 9 Evans
tube 10 Peking
tube 11 S100
tube 12 Tokyo
tube 13 Williams 82
tube 14 Corsoy79
tube 15 Vinton81
tube 16 Dassel
Louisiana Tech
BISC310
Lab 5: PCR of Parents
Name:_________________________ Date: ___________
Section ______________________
YOU MUST WEAR GLOVES DURING THIS PROCEDURE
Pea amplicons observed (8 reactions)
2
___
1. Label 8 0.2 ml PCR tubes (1 x 8 tubes w/caps) with your initials and sample number
(1-8)
___
2. Add 30 ul WATER to the green master mix tube labeled “2” (this tube already
contains 90 ul 2X TAQ master mix)
___
3. Add 10 ul of your DILUTED “L” primer to the tube with the PCR mix (with
green dye)
___
4. Add 10 ul of your DILUTED “R” primer to the tube with the PCR mix (with
green dye)
___
5. Spin down master mix tube
___
6. Add 15 ul of the green master mix from steps 2-5 into the labeled tubes
from step 1.
___
7. Add 5 ul of diluted DNA into the 16 PCR tubes as shown below ( we will add –
just let us know the tubes are ready)
tube 1
tube 2
tube 3
tube 4
tube 5
tube 6
tube 7
tube 8
Little Marvel
New Season
WSU31
M410
74SN3A
C580257A
New Era
Sundance 2
___
8. Spin down reactions
___
9. Place in PCR machine and note the location
Louisiana Tech
BISC310
Lab 5: PCR of Parents
Name:_________________________ Date: ___________
Section ______________________
YOU MUST WEAR GLOVES DURING THIS PROCEDURE
No amplicons observed (8 reactions)
3
___
1. Label 8 0.2 ml PCR tubes (1 x 8 tubes w/caps) with your initials and sample number
(1-8)
___
2. Add 30 ul WATER to the green master mix tube labeled “2” (this tube already
contains 90 ul 2X TAQ master mix)
___
3. Add 10 ul of your DILUTED “L” primer to the tube with the PCR mix (with
green dye)
___
4. Add 10 ul of your DILUTED “R” primer to the tube with the PCR mix (with
green dye)
___
5. Spin down master mix tube
___
6. Add 15 ul of the green master mix from steps 2-5 into the labeled tubes
from step 1.
___
7. Add 5 ul of diluted DNA into the 16 PCR tubes as shown below ( we will add –
just let us know the tubes are ready)
tube 1
tube 2
tube 3
tube 4
tube 5
tube 6
tube 7
tube 8
Williams
PI87623
PI567743
Soden Daizu
Little Marvel
New Season
WSU 31
New Era
___
8. Spin down reactions
___
9. Place in GRADIENT PCR machine and note the location
Louisiana Tech
Name:_________________________ Date: ___________
BISC310 Winter 083
Lab 6: PCR of Parents for documentation
Section ______________________
YOU MUST WEAR GLOVES DURING THIS PROCEDURE
No amplicons observed (4 reactions)
4
___
1. Label 4 0.2 ml PCR tube with your initials and sample number (1-4)
___
2. Add 15ul WATER to the green master mix tube (this tube already contains
45 ul 2X TAQ master mix)
___
3. Add 5 ul of your DILUTED “L” primer to the tube.with the PCR mix (with
green dye)
___
4. Add 5 ul of your DILUTED “R” primer to the tube.with the PCR mix (with
green dye)
___
5. Spin down master mix tube
___
6. Add 15 ul of the green master mix from steps 2-5 into the labelled tubes
from step 1.
___
7. Add 5 ul of diluted DNA into the 4 PCR tubes as shown below (we will add –
just let us know the tubes are ready)
tube 1
tube 2
tube 3
tube 4
84-1
4290
4895
Jackson
___
8. Spin down reactions (leave in plate, spin in plate centrifuge)
___
9. Place in PCR machine and note the location below (Which columns are your
reactions loaded in?)