Download Cloning of genes from genomic DNA Parts 4 and 5: Ligation and

Cloning of genes from genomic DNA
Parts 4 and 5: Ligation and Transformation
So far, we have isolated genomic DNA, used PCR to amplify two specific genes from the
genomic DNA, and have digested those PCR products with restriction enzymes (RE). These enzymes
should cut sites located in the PCR primers to create “sticky ends” for cloning. As we designed the
experiment properly, there are only restriction sites for the enzymes in the primer sequences and not in
the body of the PCR product. We will use these sticky ends to clone our PCR products (insert) into the
pBluescript (pKS) plasmid (vector) during the ligation and then place these recombinant plasmids
(vector + insert) into DH5α, a lab strain of E. coli, during the bacterial transformation. The bacteria
provide us with a way to isolate and generate large amounts of the recombinant plasmid.
Ligations. In addition, Tina and I (which really means Tina) have linearized the plasmid vector,
pBluescript, by cutting it with the appropriate enzymes (XbaI and HinDIII). This week, we will ligate
the cut PCR product and the cut vector together, thereby cloning our gene. The “sticky” ends (5’
overhangs) created from the restriction enzyme digestions will allow the XbaI end of the plasmid to
basepair with the XbaI end of the PCR product. The HinDIII ends will also basepair to each other.
Then DNA ligase will be able to form the phosphodiester bond between the two fragments, ligating
them together. The end result of the ligation reaction should be the formation of a pBluescript vector
containing either the eve or twist insert (PCR product). In order to tell whether the ligation was
successful, you will transform the products from your ligation reaction into E. coli and select for
bacteria containing your new plasmid.
Transformations. Transformations will be performed in much the same way as we did them at the
beginning of the semester. There are a few differences. First, we are transforming using ligation
products instead of purified plasmid. Purified plasmid transforms cells very efficiently. The ligation
products contain all sorts of mixtures of molecules and tend not to transform the cells nearly as well.
In addition, because all of our reactions (enzyme digests, ligations, etc) have some degree of
inefficiency, we can sometimes get plasmid without any inserted PCR product in the ligation mix.
Plasmid with PCR product and plasmid without any PCR product will both confer ampicillin resistance
to the transformed bacteria. In order to tell the difference between the plasmid alone and the plasmid
with our inserted PCR product you will perform blue-white screening. This selection process was
planned in advance for your experiment and is described below.
What is blue-white screening? It turns out that the pBluescript plasmid was designed with its
mulitple cloning site (MCS) in the middle of the lacZ gene. The lacZ gene, you’ll remember, encodes
β-galactosidase which is the enzyme capable of cleaving X-gal (when we include it in the plates) and
turning the colonies blue. What do you think happens if we stick a piece of DNA (say, our sim or sna
PCR product) smack dab in the middle of the lacZ gene? Usually, we disrupt the lacZ gene such that it
is unable to produce a functional β-galactosidase enzyme. Hence, our experiment will have…
1) a selection component: only those bacterial cells in the transformation that get any plasmid will
survive (ampicillin resistance) and,
2) a screening component: only those bacterial cells that received a plasmid with an insert will be
white. If they did not receive an insert, the lacZ gene will produce a functional protein and the colony
will turn blue.
Figure 2. Diagram of pKS+ plasmid, pBluescript, that we are using in lab. The multiple cloning site
(MCS) is contained at the 5’ end of the β-galactosidase gene (lacZ) and contains the restriction enzyme
(RE) site for both XbaI and HindIII used to prepare the plasmid for ligation with the sim or sna PCR
products digested with EcoRI and HindIII. Some other RE sites are shown on the plasmid along with
the ampicillin resistance gene, bla (encodes β-lactamase) and the origin of replication, rep. Enzymes which
cut pKS+ once are Acc65I 755, AdeI 222, AflIII 1153, AloI 169, ApaI 749, BamHI 689, BcgI 2562, BcuI 683, BsaAI 225, Bsp120I
749, BstXI 658, Bsu15I 725, BtgI 661, CaiI 1564, Cfr9I 695, Cfr42I 661, Eam1105I 2041, Ecl136II 653, Eco31I 2113, Eco32I 713,
Eco52I 670, EcoO109I 748, EcoRI 707, FaqI 457, GsuI 2131, Hin1I 2582, HincII 734, HindIII 719, KpnI 755, NotI 669, OliI 659,
PdiI 328, PdmI 2641, PscI 1153, PsiI 97, PstI 701, SacI 653, SalI 734, SapI 1030, ScaI 2524, SmaI 695, TatI 2524, XbaI 677, XceI
1153, XhoI 740, XmiI 734. Taken from http://www.fermentas.com/techinfo/nucleicacids/mappbluescriptiiskks.htm .
Ligation Reaction:
(Get ice)
1. Ideally the ligation reaction should have a 3:1 molar ratio of insert to vector DNA. This just means
that you want 3 times the number of insert molecules compared to the number of vector molecules in
the reaction. Given that our vector is 3 kb in length and the insert is smaller in length (to make the
math easier let’s say 1/3 the length), this means you want approximately 1:1 ratios by weight (ng) of
insert to vector DNA in the final ligation reaction. One way to think about it, is that it takes ~ 3 insert
molecules to have the same length (and thus weight) as a single vector molecule giving us the 3:1
molar ratio when the weights are equivalent.
Although we normally aim for 150 to 300 ng of vector (pBluescript) in our reaction when possible, a
successful ligation and subsequent transformation can be performed with less DNA. We will discuss in
lab how much vector and insert to use in each ligation. You will need to determine the concentration of
your insert by analyzing the gel you ran and from this calculate the number of microliters that will give
you the correct amount (ng) of insert to use. The concentration of the vector will be provided.
sim or sna Ligation (insert + vector):
? µl insert (PCR, digested and cleaned, saved from last week’s PCR clean up kit)
? µl digested pBluescript , HindIII/XbaI digested and cleaned
10 µl 2x DNA ligase buffer
? µl dH20
20 µl total volume
We will set up one control ligation per lab day (volunteer group????):
Control Ligation (vector only):
? µl digested pBluescript , HindIII/EcoRI digested and cleaned (use same amount of vector as above)
10 µl 2x DNA ligase buffer
? µl dH20
20 µl total volume
2. Add 1 µl of DNA Ligase to your tube(s). Flick the tube to mix and then pulse-spin to bring all of
your sample down to the bottom of your tube.
3. Incubate at room temperature for 5 minutes.
4. Store your sample on ice while getting ready to transform DH5α competent cells.
Bacterial Transformation:
You will be transforming some of your ligation mixture into the DH5α strain to identify a plasmid
containing the insert using blue/white screening. These cells have been specially prepared to have a
high efficiency of transformation, thus increasing the number of colonies that you get from the
transformation reaction of the ligation products. What color should a colony have if it has been
transformed by plasmid containing the insert? The control ligation will also be transformed to see what
background is obtained from the vector ligating back on itself. The vector has been treated with a
phosphatese to minimize this problem but it can still occur at a low frequency. Do you expect a lot of
transformed cells from this ligation? Why? What color would these colonies be on LB amp IPTG Xgal media?
1. Thaw the DH5α competent cells on ice.
2. Place a sterile 5 ml polypropylene snap cap tubes on ice. Add 50 µl of competent cells and 2 µl of
your ligation reaction. Gently swirl to mix.
3. Incubate on ice for 30 minutes.
4. Heat shock the cells at 42°C for 30 seconds.
5. Immediately place back on ice for 5 minutes.
6. Add 950 µl LB broth and incubate at 37°C for 20-60 minutes, mix occasionally. The timing is
dependent on whatever else you may be doing in lab, anytime within this range will work perfectly
fine.
7. Plate 100 µl on one LB amp IPTG X-gal plate and 50 µl on another LB amp IPTG X-gal plate.
Incubate the plates upside down at 37°C overnight. Label your plates well so it’s clear what is in
each plate.
8. The following morning, your plates will be taken out and stored at 4°C (this will be done for you).
As a side note, if the plates are left in the incubator too long you will get satellite colonies (bacteria
with no plasmid) growing around your transformed bacterial colonies in areas where the ampicillin
has been depleted. This makes it especially difficult to pick colonies. Next week you will pick 2
appropriate colonies, each containing a unique recombinant plasmid, and grow them up in a liquid
media the day before lab in order to isolate the plasmids during the lab. Directions for starting
your overnight cultures are provided in the next handout.
*** Note, this semester we will streak out colonies (4-8) individually on a LB amp IPTG X-gal plate 2
days before the lab and use 2 of those colonies the next day (1 day before lab) to start our overnight
cultures from instead of going directly from our transformation plates. This will be explained in more
detail in lab.