Genetics Lab Write-up Outline Title: Cloning of the Drosophila melanogaster Tran

Genetics Lab Write-up Outline
Title:
Cloning of the Drosophila melanogaster Transformer Gene
Abstract: 1 paragraph:
-Summary of Drosophila melanogaster as a model organism and the function of the
Transformer gene protein.
-Goal: to amplify and clone and sequence a portion of the Transformer gene for gene
expression studies
-Summary of methods:
genomic DNA isolation, PCR, ligation into vector, transformation into cells,
plasmid DNA isolation, plasmid sequencing
-Summary of results:
successful amplification and cloning the Transformer gene
Introduction ~1 page, at least 3 references:
-Background on Drosophila melanogaster as an important model organism (ref)
-Background on Transformer role in cell biology/role, gene expression studies (ref)
Materials and Methods ~2 pages, at least 4 references:
-genomic DNA isolation from adult Drosophila melanogaster (ref)
-primer sequences (should give gDNA PCR product of 399bp)
dmTra-F Forward TGAAAATGGATGCCGACAGC
dmTra-R Reverse ACCTCGTCTGCAAAGTACGG
(Integrated DNA technologies, Carlsbad CA)
-PCR (ref) 56C annealing temp, 30 cycles
-ligation into vector (details? vector, Invitrogen, Oceanside CA)
-transformation into competent cells (details? cells, Fisher Scientific), cell culture
and blue/white screening (ref)
-plasmid DNA isolation (ref)
-restriction digest with EcoRI
-restriction digest gel
-sequenced plasmid product (UC Berkeley Sequencing Facility)
Results 2-3 pages:
Written summary of following results:
-DNA 230/ 260/280 values, discuss concentration of DNA, any contamination (in
notebook)
Figure 1 -PCR gel picture with legend (your picture)
Figure 2 -Bacteria plate picture with legend (your picture)
Figure 3 -plasmid gel picture with legend (in notebook or Canvas)
Figure 4 -Ensembl BLAST Search Alignment (screenshots from web)
Here is my lab notebook with all the procedures and data you require.
BIOS 332 Genetics
Lab #2 Isolation of Genomic DNA from Drosophila melanogaster
Please wear pants in addition to closed toed shoes to this lab! I highly
recommend not wearing contact lenses as well.
Background:
We will be isolating genomic DNA from adult Drosophila to use as the starting material
for our cloning project. In order to isolate the DNA from the flies, we will first have to
lyse (break open) cells. This will expose all the cellular contents to the same solution and
the DNA, RNA, proteins, sugars and lipids will all be in the mix. We will lyse the cells in
a detergent called sodium dodecylsulfate (SDS) that will help to disrupt the lipid
membranes as well as denature (unfold) and inactivate all the proteins in the sample.
Subsequent steps will purify the nucleic acid from the sample and then selectively
remove any RNA leaving a pure genomic DNA (gDNA) sample.
SDS
We will first isolate
all the nucleic acid
(DNA and RNA)
away from the other
cellular components.
Briefly, we will lyse the cells in the tissue by homogenization in an SDS buffer and then
incubate with proteinase K to digest the cellular proteins into short peptides. We will then
separate the nucleic acids by phenol/chloroform extraction. Phenol
is a protein solvent: the hydrophobic ring will penetrate the
hydrophobic core of globular proteins and denature them, and the
hydroxyl group will solubilize hydrophilic side chains in the
peptide. Phenol is only partially soluble in water and so will form a
separate phase when added to aqueous solutions. Proteins, lipids
from the cell lysate and SDS will partition into this layer and in a
whitish material between the two phases. You will want to avoid
Phenol
this material. When you are working with phenol you will want
to wear gloves and safety glasses. You don’t want to denature the proteins on your
skin and in your eyes! We will also add chloroform to our organic phase. Chloroform is
a heavy organic molecule that will help to make a distinct interface between the aqueous
phase and the organic phase. The stock bottle of phenol/chloroform will be overlaid with
an aqueous layer so be sure to take the lower phase. In our sample, the nucleic acid will
remain solubilized in the aqueous phase, and we can remove this to a new tube. We will
concentrate the nucleic acid by precipitating it in a salt/ethanol solution and resuspend it
in a smaller volume of water. We will then selectively digest away the RNA with RNAse
enzymes leaving behind a pure gDNA sample. One last important point: Your hands
are covered with enzymes that could potentially degrade the DNA so you need to wear
gloves throughout the procedure.
Materials:
Adult flies
Lysis buffer (SDS buffer)
Microcentrifuge tubes and plastic pestles
10 mg/ml Proteinase K
1:1 phenol:chloroform solution
Gloves
Pellet paint
3M Sodium Acetate
Ethanol (95% and 75%)
RNAse and 10X RNAse buffer
RNase/DNase-free water
Microcentrifuge, 37º C and 42º C water baths.
Crushed ice
Procedures:
A. Nucleic acid isolation:
1. Add 200 μL of lysis buffer to a blue 1.5 mL centrifuge tube. Add 15-20 flies to
the tube.
2. Insert the plastic pestle and grind the flies against the bottom and sides of the tube
a dozen or more times.
3. Add another 300 μl of lysis buffer with 12μL of 10 mg/mL proteinase K solution
(final concentration ~250 μg/mL). Mix by inverting the tube several times.
(Accidentally added 350 ml so now total Volume is 550 ml)
4. Incubate for 30 minutes at 42º C.
5. Add equal volume (0.5 mL) of phenol/chloroform to your tube and mix by
inverting the tube several times. Centrifuge for 3 min at full speed and carefully
remove 450 μL of the aqueous phase (upper layer) with a P1000 pipetteman, being
careful not to take any of the white material at the interface. (It’s better to not get all
450 μl than to contaminate with the white interface.)
6. Now repeating step 5 with your new tube making sure to add equal volume of
phenol/chloroform (450 μL) this time.. You should have very little white at the
interface now. Remove 400 μL of the top aqueous phase to a new tube.
7. Add 100 μl of Nuclease Free water to bring the volume up to 500 μL.
B. DNA isolation
1. To your aqueous phase add 50 μL of 3M sodium acetate and 1mL of 95-100%
ethanol and 1 μL of pellet paint (I will give this to you personally) Mix well by
inversion and put your tube on ice for 15 min. (This will cause the nucleic acid to
precipitate out of solution.)
2. Centrifuge full speed at 4º C for 15 min. You should see a small red/pink spot on
the side of the tube near the bottom indicating where your nucleic acid pellet is
located. Pour the ethanol off into a new tube (your pellet should remain stuck to the
tube). Keep your eye on the pellet so you don’t lose it!
3. Add 100 μL of 70% ethanol and re-centrifuge for 3 minutes. (This will leach out
some of the sodium from your pellet). Pour the ethanol off again, making sure you
don’t lose the pellet. Dab any droplets with a twisted Kimwipe, being sure not to
touch the pellet.
4. Let the pellet air dry for 5 min with the tube lying on its side on the bench.
5. Add 44 μL of Nuclease-free water to the tube and resuspend the pellet by
pipetting up and down until it has completely dissolved.
STOP: Do not proceed unless you have at least 40 minutes left in the lab session.
6. To the tube add 6μL of the “RNase Cocktail”:
5 uL 10X RNAse buffer
1 uL RNAse (DNAse-free)
7. Incubate at 37 º C for 20 min. (This will digest away all RNA leaving only DNA)
8. Bring up the volume by adding 150 μL of nuclease-free water.
9. Extract one time with 200 μL of phenol/chloroform and move 180-200 μL of the
aqueous phase to a new tube. (To get rid of the remaining proteins)
10. Add 20 μL of 3M sodium acetate and 440 μL of 95-100% ethanol, mix by
inverting and store in the freezer until next lab. Make sure your tube is labeled
clearly!
Congratulations! You should now have a pure genomic DNA sample. We will check the
quality and quantity of our samples next week. In your discussion please comment on any
mistakes you made, alterations to the protocol that were necessary and any observations
you made.
Added an extra 50 ml of Lysis Buffer, will have to pippette more material out or add less material to fix volume.
added 550 phenol chlorform
removed 500
added 400 phenol chloroform
removed 400
added 100 ml nuclease free water
added 50 ml and 1000ml
drained
performed experiment normally from here since volume resets after draining
Discussion:
Did not wait 5 min for ethanol to dry off pellete, proceedeed with experiment as normal
BIOS 332 Genetics
Lab #3 Analysis of your DNA sample, PCR
Background:
Before we move on to PCR using our genomic DNA as a template, we need to have an
idea of the amount and purity of our DNA sample. Typically, this is done with an
ultraviolet (UV) spectrophotometer. Nucleic acids (both DNA and RNA) absorb UV light
maximally at 260 nm. The amount of DNA will be proportional to the absorbance units
(AU) at 260 nm (AU260). A DNA solution with an AU260 of 1.0 has a concentration of
0.05 μL / μL. So you could calculate your samples concentration by the following
formula:
AU260 * 0.05mg/ml * dilution factor = DNA conc. in mg/ml.
Typically, you would also take a reading at 280 nm (also in the ultraviolet range).
Nucleic acids only partially absorb this wavelength, but proteins have their peak
absorbance at this wavelength. By calculating the 260/280 ratio you could estimate the
purity of the sample. Pure DNA would give a ratio of ~1.8. Anything less than this would
indicate that your sample was contaminated with protein or other organic molecules that
absorb in the 280 range. You will also calculate the 260/230 ratio. Pure DNA should give
you a ratio of 2.0-2.2. Anything less that this would indicate contamination with organic
molecules that absorb primarily in the 230 range (like phenol).
We will also use the “Qubit” dye chemistry and fluorometer to calculate the DNA and
protein concentrations in our sample. This procedure uses chemical dyes that specifically
bind to DNA, RNA or proteins and the absorbance of the dyes at visible wavelengths can
be used to determine concentrations.
Once we have determined the quantity and purity of your genomic DNA sample, we will
use it as a template for PCR with primers we design to amplify our DNA sequence. PCR
is a simple procedure: You will add template DNA (genomic DNA in our case), the
primer pair(s), deoxynucleotides with an equal mix of the four bases (dNTPs), DNA
polymerase (Taq) and a magnesium buffer solution to ensure the enzyme will be active.
The thermocycler machine will do the rest. It will cycle between 98° C (denaturing) to a
primer-specific annealing temperature. We will set this temperature based on the size and
estimated melting temp of the primers. Typically, this is between 50-60° C. If this temp is
too high no primers will anneal. If it is too low the primers will anneal to sequences that
are not fully complementary and we will get non-specific products. Lastly, the machine
will cycle up to the elongation temp (68-72° C) where the polymerase has its optimal
activity level. I will set the machine to cycle through this series of temps 25-30 times.
Procedure:
A. Finish Ethanol precipitation:
1. Spin down your sample from last lab (100% ethanol) at full speed for 15 min to pellet
your RNA. You may or may not see a pellet.
2. Pour off the 100% ethanol and add 100 μL of 70-75% ethanol. Spin for another 3 min
at full speed.
3. Very carefully pour off the 75% ethanol into a catch tube (be sure not to lose the
pellet). Dab off any droplets in the tube with your Kim wipe probe and let the pellet dry
for about 5 min laying on it’s side on the bench at room temp.
4. Re-suspend the pellet in 45 μL of nuclease free water.
B. Nanodrop Spectrophotometry.
1. In small groups, your instructor will escort you to LIM 279. Transfer 1 μL of your
DNA solution to the nanodrop pedestal and lower the arm.
2. Record your 260/280, 260/230 and your concentration. You may also have an adjusted
final concentration. Take a picture and upload it to your benchling entry.
4. Lastly, calculate what volume of your sample you would need to have 500ng of DNA.
260/280: 2.05
260/230: 1.31
ng/ul: 297.7
1.68ng
C. Qubit Dye Fluorometry.
1. Pipet 198 μL of “Protein Working Solution” into an assay tube.
2. Add 2 μL of your sample, vortex for 2 seconds and incubate at room temp for 15 min.
3. Pipet 198 μL of “DNA Working Solution” into a second assay tube.
4. Add 2 μL of your sample, vortex and incubate for 2 minutes.
5. Once your incubations are complete, take your tubes to the “Qubit” spectrophotometer
to read the concentration of your sample at (I will help you run the spectrophotometer)
6. Record the protein and DNA concentrations in your notebook.
protein: 62.7 0.627. ug/ml
dna: 41.7, 0.417. ug/ml
D. Set up PCR reactions
1. Each group should set up three PCR reactions:
1. DNA from student #1, Experimental Primers
2. DNA from student #2, Experimental Primers
3. Negative control (no template DNA), Experimental Primers
(Your instructor will set up two positive controls: one with a verified template and
experimental primers, another with verified template and verified primers)
2. Mix PCR reactions in a new (very small) PCR tube:
-25 μL of the PCR Master Mix
-5 μL of the primer mix (both forward and reverse primers included)
-____1.68_μL of template gDNA (for ~500ng) (or 1 μL) 2
– __23__μL of nuclease free H2O (so final volume is 50 μL)
3. Label your tubes and place them in the labeled rack on ice.
4. Your instructor will start the thermocycler once everyone has finished.
Discussion:
Everything went smoothly, no issues during the procecdure.
Lab #4 PCR, Agarose Gel Electrophoresis
Procedure:
1. Make 2% agarose gel (1g agarose in 50mL of 1X TAE) in a 200mL
Erlenmeyer flask with a kimwipe plug
-microwave for 30 seconds and then swirl then repeat 2 more times (1min, 30sec
total)
-Let cool on the benchtop for ~5 min
-Add 2 ul of 10 mg/ml Ethidum Bromide and swirl. (in hood: wear gloves, glasses
and be very careful!)
2. Pour ~40mL of the molten gel into the electrophoresis casting tray. Make sure
rubber seals are well seated and the comb is properly positioned. Let cool 15-
20min.
3. When the gel is cool and solid to the touch, remove combs and re-orient you
gel casting tray so wells are near the anode (negative pole) and then pour in 1X
TAE buffer until the gel is fully submerged under the buffer.
4. Add 3 μl loading dye to 10 μl of your PCR sample and load all 13 μl into a
well of your gel. Your instructor will load 8μL of the DNA ladder.
5. Put on the lid, making sure the DNA in the wells is going to move through the
gel toward the cathode (positive (red) pole). Connect the wires to the power
supply and turn the power to 100V.
6. Look for bubbles (good) or sparks (bad) coming off the electrodes. This
indicates that the current is running, and no dangerous charge is building up.
7. Watch the dye migrate. You want the yellow dye to run one half the length of
the gel.
8. Look at your gel under the UV transilluminator. Ideally, we will see a single
band in each experimental lane and nothing in the negative control lane.
exACTGene 1Kb Plus ladder:
Discussion:
Everything went accordingly as planned
gel also proved good results
Lab #5 Fly Assessment
1. Assess health of the vial
2. Verify the phenotypes
3. Clear the vial of all adults (fly morgue)
4. Wait 2 hours and collect virgin females
5. Setup genetic cross in new tube 3-5 virgin females bb normal with 5-7 vestigial males
Disucssion:
Flies were well
Lab #6 TOPO Cloning and Bacterial Transformation
Background:
Previously we successfully amplified the fragment of the Transformer gene by PCR.
Today, we will clone the PCR product into a plasmid vector and transform E. coli cells
with our recombinant vectors. We will use E. coli cells that have been made competent
by a supply company. These cells have been specially treated with high salt concentration
or an electrical current to permeabilize the membrane, making their uptake of
extracellular DNA more efficient when we heat shock them. Only some of the cells will
take up our plasmid during the heat shock, so we will need to screen the bacterial so only
those that took up the DNA will grow. We will do this by plating the cells on an agar
plate that has the antibiotic ampicillin in it. Only the cells that have gained the amp
resistance gene by taking up the vector should be able to grow; all other cells will die off.
Additionally, we will add the chemical X-gal to the plates in order to perform blue/white
screening for bacterial colonies that have a vector that contains an insert in the multiple
cloning site (MCS). Our insert will have disrupted the function of the LacZ (ß-
galactosidase) gene in the MCS making it unable to cleave X-gal. Colonies with an insert
will appear white, while colonies with a functional LacZ gene will be blue.
Next time we will pick several white colonies and grow them in larger quantities in a
liquid culture and isolate plasmid DNA. We will perform a restriction digest to check for
the presence of an insert that is the same size as the PCR product we intended to clone in.
We will choose one of these DNA samples to send off to be sequenced.
Materials:
LB-amp plates (100μg/mL ampicillin)
Chemically competent E. coli (Mach-1 chemically competent cells)
S.O.C. Medium
42°C water bath
37°C incubator and shaker
Cell spreaders and Isopropanol
40mg/mL X-Gal
50mg/mL Ampicillin or Kanamycin
Procedure:
Part A: TOPO cloning (do not start until instructed to do so!)
1. Centrifuge the pCR BLUNT II-TOPO vector tube to collect the liquid in the
bottom of the vial.
2. Determine volume of PCR product needed for a 1:1 vector:insert ratio (best for
inserts 400-700bp). For a typical PCR reaction this should be 2 μl.
3. Set up your ligation reaction in a PCR tube as follows:
Fresh PCR product 2μl
Salt Solution 1μl
Sterile water 2μl
Vector (25ng/ul) 1μl
Total volume 6μl
4. Incubate the ligation at room temp for 5 minutes. Continue directly to part B.
Part B: Transform Cells
1. Take one 50 μL vial of competent cells and thaw on ice (~10 minutes)
2. Pipette 2 μL of your ligation reaction directly into the vial of competent cells and
mix by stirring gently with the pipette tip.
4. Let the cells sit on ice for 15 min. Return your ligations to the ice container in the
front.
5. Heat shock the cells at 42° C for 30 seconds exactly and immediately put them
back on ice.
6. Add 250 μL of room temp. S.O.C. medium and incubate at 37° C with shaking for
40 min.
7. While your cells are shaking, spread 40 μL of 40mg/ml X-Gal and 150 μL of
1mg/mL kanamycin on two pre-warmed LB plates. Invert them, label them on the
bottom with your group’s initials and “50 μL” on one and “100 μL” on the other.
Return them to the 37° C incubator.
8. Spread 50 μL and 100 μL of your transformed cells on to the respective plates.
Invert and incubate overnight at 37° C. I will transfer your plates to 4° C tomorrow
morning. The cells will be fine on the plates until we pick colonies next week
Discussion:
Everything went accordingly.
Lab #7
Plasmid DNA minipreps.
Background:
Last night, I picked six white colonies of bacteria that should contain a plasmid that has
your PCR product inserted into the multiple cloning site. Each colony is the result of a
single bacterial cell transformed with a single plasmid from the ligation reaction. I
inoculated 3ml cultures and incubated them at 37 C with shaking overnight (~18 hours).
Each tube should contain copious numbers of bacterial cells each with multiple copies of
the ligated vector.
Today you will isolate the plasmid DNA from each of these cultures in a method called
plasmid minipreparations or “minipreps” for short. Plasmid DNA preps are named
according to the DNA yield that is expected from a given volume of cultured cells.
Minipreps cultures of 3-5ml typically yield 20-30 μg of plasmid DNA depending on the
strain of bacteria used (different bacteria average different numbers of plasmids per cell).
Other prep varieties are:
Midipreps use 15-25 ml of media and the expected DNA yield is 100-350μg.
Maxipreps use 100-200 ml of media and the expected DNA yield is 500-850μg.
Megapreps use 500 ml – 2.5 L of media and the expected DNA yield is 1.5-2.5 mg.
Gigapreps use 2.5-5 L of media and the expected DNA yield is 7.5-10 mg.
Procedure:
A. Document your LB-Amp, X-Gal Plate
-Take a picture of your best plate for your notebook and write-up.
B. Plasmid “miniprep” Isolation
1. Transfer 1.5 ml of each of your liquid cultures to labeled microfuge tubes. Save the
remaining 1.5mL of your culture in the test tubes.
2. Spin your tubes at full speed for 30sec to pellet the cells. Pour off all the liquid into
one of the labeled waste beakers. Add the remaining 1.5 mL of liquid culture from the
test tubes into the same microfuge tube. Spin again for 30sec and pour off the rest of the
liquid.
3. Resuspend the pellets in 100 μL of Resuspension Solution (Solution I) by pipetting up
and down and then vortexing. Be very thorough, and make sure there is no pellet left.
You may want to hold your tubes up to the light to ensure you sample is homogeneous.
4. Add 200 μL of the Cell Lysis Solution (Solution II) and GENTLY (no vortexing) rock
the tubes back and forth while holding them horizontally. This solution will disrupt the
bacterial cell wall and plasma membrane allowing the cellular contents (including your
plasmid DNA) to spill out into the solution. Let the tubes rock on the platform rotator for
~5 min. The solution will become thick but transparent with clumps of cell debris visible.
In bacterial cells, the genome is tethered to the plasma membrane and if we are gentle it
will remain with the cellular debris and our miniprep sample will be pure plasmid DNA.
5. Add 150 μL of Neutralization Solution (Solution III). Again, mix gently by rocking
the tube sideways. The KOAc will tend to sink to the bottom of the tube, so be sure your
mixing is thorough, but gentle. Mix until the solution loses its viscosity. Let the tubes
rock on the platform rotator for at least 10 min.
6. Centrifuge at full speed for 2 minutes to pellet the cell debris. Pour the supernatant
into a new, labeled tube. You should have about 400 μL.
7. Add equal volume (~400 μL) of 1:1 phenol/chloroform. Mix gently by inverting the
tubes a couple of times at first, then vortex for a couple of seconds. Centrifuge for 3 min
to separate the phases. Remove ~350 μL the aqueous (upper) phase to new, labeled
tubes, making sure you do not suck up any phenol or material that may be sitting at the
interface of the two phases. It is better to leave some aqueous phase behind, then
contaminate with phenol or stuff at the interface! Put the leftover tube in the phenol waste
rack.
8. Precipitate the DNA by adding 2 volumes (~700 μL) of 95-100% EtOH to the tubes
and invert them several times. Let them sit in on ice (or in the refrigerator) for at least 15
min.
9. Spin the tubes at full speed for 10 min and pour off the supernatant. Orient your
microfuge tubes so the hinge is toward the outside edge of the rotor. You should see a
white pellet on the side of the tube. This is cellular RNA and your plasmid DNA.
1O. Wash the pellet by adding 200 μL of 75% EtOH. Do not try to resuspend the pellet.
Instead, immediately spin again at full speed for 2 minutes.
11. Pour off the 75% EtOH, being careful not to dislodge the pellet. Dry and then
resuspend the pellet in 20 μL of TE solution.
Store your miniprep DNA samples in your rack in the refrigerator until next week, when
we will digest away the cellular RNA, perform a restriction digest on the DNA and run
an agarose gel to check the size of the insert in each of the samples.
Materials:
1.5ml microfuge tubes
microfuge
Resuspension Solution (Solution I)
Vortex mixer, platform rotator
Cell Lysis Solution (Solution II)
Neutralization Solution (Solution III)
Buffered Phenol/Chloroform
100% or 95% EtOH
TE buffer
Appendix:
Solutions used:
Resuspension Solution (Solution 1)
25mM Tris-HCl, pH 8
10mM EDTA
15% Sucrose
Lysis Solution (Solution II)
0.2M NaOH
1% SDS
Neutralization Solution (Solution III)
3M Potassium acetate, pH 4.8 (pH with acetic acid)
RNase Solution (TE plus RNase)
50mM Tris-HCl ph 8
1mM EDTA
Add RNase A, T1 at 10 μg /ml and 10units/μL to digest
Discussion:
Everything went accordingly except samples 1 and 2 were vortexed at the start when they shjouldnt have been.
Sample 3 remains pure and the best viable option
Lab #8 Plasmid Restriction Digests
Background:
Today we will finish the isolation of our plasmid DNA by digesting away all of the
remaining cellular RNA that remains in our sample. Remember, DNA (genomic and
plasmid) and RNA have essentially the same chemical properties and so will be isolated
alongside each other in a phenol:chloroform extraction and alcohol precipitation. We
will use the Ribonucleases RNase A and RNase T1 to selectively digest the RNA, leaving
the plasmid DNA intact. Next we will use the restriction enzyme EcoRI to digest your
plasmid DNA and we will run the fragments on an agarose gel in order to identify the
size of your insert. Your insert will vary in size depending on the primers that were used
in the PCR reaction. I will do a final purification of the successful samples from this
week and send them to be sequenced.
Materials:
-Enzymes: EcoRI, RNAse A, RNase T1
-10X Restriction Enzyme Buffer
-Sterile water
-37° C water bath
-Agarose
-1X TBE or TAE buffer
-Submarine gel electrophoresis apparatus
-10 mg/ml Ethidum Bromide
-DNA loading dye
-DNA ladder
-UV Transilluminator
Discussion:
No issues, lab proceeded as expected.
Procedure:
A. Remove 2 μL from one of your plasmid DNA samples and put in a new tube to
use as a “no RNAse” control sample
B. RNA digestion:
1. Add 4μL of RNase cocktail to each of your plasmid samples and incubate in a
37° C water bath for 15 min.
C. EcoRI Restriction Digest and Gel Electrophoresis
1. While your RNase digest is incubating set up the following in three new tubes:
10 μL sterile water
2 μL 10X Restriction Enzyme Buffer
2 μL EcoRI Solution
2. Once the RNase digestion of your plasmid DNA is complete, add 6 μL to the
restriction enzyme reaction.
3. The EcoRI contains glycerol (which will make it settle to the bottom of the
tube) so be sure to mix the components well by pipetting up and down.
4. Incubate the tubes for at least 30 min in a 37° C water bath.
5. While your digests are incubating, pour a 2% agarose gel:
-1g in 50mL of 1X TBE or TAE buffer
-Let cool for ~5 min
-Add 2 μL of 10 mg/ml Ethidum Bromide and swirl. (wear gloves and
goggles and be very careful!)
-pour into the electrophoresis casting tray
6. At the end of your incubation add 6 μL of the 6X DNA loading dye to each of
the tubes and mix well
7. Load 10 μL of each of your samples on your gel. Your instructor will load one
lane with 10 μL of the DNA ladder.
8. Run your gels at ~100V until the dye has migrated at least one quarter the
length of the gel. Look at your gel on the transilluminator and photograph for
your notebook. We are hoping to see a band at that corresponds to the PCR size
and a second band at ~4000bp that is the plasmid.
We will choose one or two of our best samples to send out for sequencing later
this week
Discussion:
PCR insertion was not successful, sample 2 only showed a smeared DNA sample.
Lab #9
Fruit Fly Maintenance and review
Discussion:
75 Pol2 example sequence
GAATTCGCCCTTAGGGCGGCGAGGACATGGATCTCACCAAGGAGAACCAGCAGCCGGATCCAAACAAAAAGCCCGGCCACGGCGGTTGCGGTCACTACCACCGTCGAAGGGCGAATTC
Learned how to effectively use Finch.tv and cleanup dna sequence.
Lab #10 Sequence analysis of Drosophila Transformer clone.
Background:
The sequencing facility at UC Berkeley has run our samples on their Applied Biosystems
96 capillary 3730xl DNA Analyzer. The lab used use a primer specific to the SP6
sequence in the vector and used fluorescent ddNTP’s to perform the dideoxy sequencing
reaction. Instead of using a gel to run the samples the 3730xl machine used polymerized
acrylamide in tiny capillary tubing. The machine collects the sequence data and they have
e-mailed us a digital file of the chromatogram. The files are named “28-tra-
1_D04_026.ab1, 29-tra-2_E04_024.ab1”, and so on. We will need to clean the data up a
bit and then you will do some simple analysis using free software tools available online.
As you go you should answer the questions in your notebook.
Materials:
Computer wireless access
Mek&Tosj 4Peaks DNA chromatogram (Mac) viewing software
Download from : https://nucleobytes.com/4peaks/index.html
OR
FinchTV DNA chromatogram viewing software (Mac or PC)
Download from: https://digitalworldbiology.com/FinchTV
Map of the vector (included in this protocol)
Transformer gene primer sequences (included in this protocol)
Procedure:
Part A: Sequence clean up:
1. Download all four .ab1 files from Canvas and save it to the desktop.
2. Open the 4Peaks or Finch TV software and open your file from the “File” dropdown
menu.
3. You should be looking at your sequence displayed in 4 color-coded peaks (see below).
The programs will do some auto-analysis on the data and will be displaying the base
sequence above each peak in the chromatogram.
4. The first thing we will do is look for any Ns in the first ~500 bases and try to
determine what the base is by looking at the chromatogram. Don’t worry about the first
10 or so bases: these peaks represent unincorporated fluorescent nucleotides and
fragments that were too small for the machine to resolve. If there are large numbers of Ns
past ~600bp go ahead and leave them, we are going to delete this sequence anyway. You
can change the base from an N to the appropriate base by double clicking the letter and
typing the correct letter.
5. “Clean up” the sequence by fixing as many incorrect bases as you can. Look for
miscalled bases, inserted bases or missed peaks. Also look for minor peaks that may
show up beneath the major peak that the machine has called automatically. Ask for help
if you run into any problematic sequences. If you really cannot determine the correct
base, just leave the “N”.
6. Once you have “clean” sequence data, we need to delete the vector sequence before we
can analyze our sequence. Use the search tool to find the EcoRI site (GAATTC) that
should precede your inserted sequence. Compare the flanking sequence to the vector
map below and verify that this is the vector sequence you suspect.
7. Select the entire identified vector sequence and delete it by using “Edit:Delete
Selection” or right click (control click) and you can choose “Delete” from the drop menu.
8. Now your sequence should begin with the sequence of one of your two primers. The
PCR product could have inserted in either direction, so you will have to determine what
orientation it is in. Your sequence may be reading as the inverse complement of what the
map below shows, so be careful.
Part B: Sequence Analysis:
1. Copy the “clean” sequence to the clipboard by using “Edit: Copy Sequence” and paste
it into benchling.
2. Let’s take each of our fragments and search the drosophila melanogaster genome with
to verify that it really is a portion of the Transformer like we think it is. Use a web
browser to navigate to the Flybase homepage and click on the “BLAST” box on the left.
Paste your sequence in the entry field and start a nucleotide (blastn NT->NT) search
choosing the Drosophila melanogaster genome. When the search is done it will display a
graphical representation of the matching sequences it found in the genome. How many
results do you get? What chromosome are these results on? How many nucleotides match
the genome? What is the score, E-value and % identity of each result?
3. Scroll down to the alignment view of the best result. On your best alignment click on
the “JBrowse” link to show the region in detail. Is this the location of the Er degradation enhancer
gene? What is the genomic location? Repeat this for all four of our sequences.
4. Once you have found which sequences are the Er degradation enhancer gene, Identify how many
introns and exons this gene has and note what region of the gene we have amplified.
Zoom in or out in order to get a screenshot of the genomic region.
5. Now that we are sure we have cloned part of the Er degradation enhancer gene, click on
“alignment” and take a screenshot. Look closely at the alignment and note any
differences between our sequence (the “query”) and the genome. Paste your screenshot
into benchling.
6. You should now have all the data and images to complete your lab write-up. Take
some time to review the lab write up criteria, rubric and sample write-up from previous
years. Contact your instructor if you need help with your write-up.
Appendix:
Drosphila Transformer (dmTra) Primers:
dmTra-F Forward TGAAAATGGATGCCGACAGC
dmTra-R Reverse ACCTCGTCTGCAAAGTACGG
Results and Disccusion:
30-tra-3 F04_022.ab1 Sequence:
GAATTCGCCCTTACCTCGTCTGCAAAGTACGGTACCGAGGAGTGACGCCATCACGTGGCGACATGGACGACATCCTTGGAAAGTGAGTTCAAAAACCTAAGGTTTGGAGGGATTACTAACATTGTTCTTATTTTTAGTTTCTCCATGACTCTGGTGGATACTCTGGACACTCTGGTGCTCCTGGGCGATTTCACAGAGTTTGAGCACGCGGTGAAGCTAGTTATCCGCGACGTTCAATTCGACAGCGACATTATTGTGTCGGTGTTTGAGACGAACATTCGAATGGTCGGTGGCTTGCTGTCGGCATCCATTTTCAAAGGGCGAATTC
1 result
2L
284 nucleotides match up
score 551.587 bits
E-value 4.02225e-156
%identity 99.3%
Yes it is the location
Location is 13193556-13193840
Our seuqence was only 2 base pairs off of the genome on Flybase. We had 99% accuracy in cloning our gene.
2 introns
4 exons