Transcription PS Answer Key – Bioreg 2004.
Problem 1
1. You are working with a novel species of bacteria. Let’s call it Bac. You want to isolate a new
RNAPolB type activity from Bac. How do you assay RNAPolB activity? What do you need to know?
Why might this be tricky, given that this is a novel species of bacteria. [6 points]
Need to know the identity of a normal coding gene or operon. Would use a plasmid containing the
promoter from this gene (or just a PCR product or something). Would first need to identify any cofactors necessary for transcription (e.g. transcriptional activators etc.). Then would do a TCA
precipitation to look for incorporated NTPs. Maybe run the NTPs on a gel. Ultimately, check that the
product is the transcript of the gene you chose. (Could use a reporter gene in an in vitro system to
facilitate this).
i)
ii)
2. You want to identify some key residues in the active site of the RNAPolB you find. Name two
methods to do this. (Pubmed search: Keyword Mustaev) [6 points]
Any kind of cross linking approach (cross link RNAPol to DNA template or to RNA). Caveat: Shows that
you are in the RNA channel but not exactly the active site.
Iron EDTA approach. Basically IRON at active site makes free radicals and causes local cleavage of
RNA or protein. Better evidence for active site.
3. You want to determine the core DNA recognition sequence for this RNAPolB. How would you do this
using a) genetics and b) biochemistry. [6 points]
Genetics: Replace regions of the promoter sequence with linker polyAs or something or make point
mutations around the putatative sites. Check reporter construct expression. Could sequence a few genes
around start site and use sequence alignments to guide you.
Biochemistry: DNAseI footprinting. Gel shifts. Small molecules. SELEX
4. You purify a transcriptional activator that enhances transcription from the Bac promoter you are
studying. You want to know if it increases the rate of transcription from gene X by a) Recruiting
RNAPolB to the promoter more efficienty (increasing KB) or b) Catalysing a rate limiting step in the
transition from the closed to the open complex. How would you distinguish between these possibilities?
[6 points]
This is essentialy the kinetic experiment described in your notes where youadd different amounts of
dinucleotide to define a Tau value for various [RNAPol]. Extrapolating [RNAPol] to infinity gives you
values for Kb and kf.
5. You want to study the regulation of elongation of RNAPolB on geneX. You decide to look for
evidence of pausing. You decide to use your technical wizardry to devise a method for looking at the rate
of elongation of a single molecule of RNAPolB in real time. How do you do this? [6 points]
DNA attached to fluorescent bead. RNAPolB imobilise by an optical trap and covalent linkage to another
imobile bead. Check the rate at which the bead moves by microscopy. Check to see if it pauses. Repeat
a hundred times.
6. It does pause in vitro! You decide to check if this happens in vivo. To do this you do the following.
You make cDNA probes corresponding to regions of gene X as in fig A.
FigA
Gene X
5’
3’
cDNA Probes
Probe:
A
B
C
D
E
F
G
You determine that you can permeabilise your Bac cells and allow transcription to continue normally as
long as your salt concentration is as in vivo. Raising the salt concentration rescues pausing nonspecifically i.e. when you raise the salt concentration, any Pol that was pause resumes transcription. So,
this is how you do your experiment (this is analogous to the Transcriptional Runoff experiments people
use a lot in the Eukariotic field):
1.
You allow your permeabilised Bac to transcribe under normal salt conditions such that, in a huge
population of cells, RNAPol is transcribing, initiating, pausing, terminating etc. as it would normally –
you have reached a steady state situation.
2.
You then raise the salt concentration so that all molecules of RNAPol are able to transcribe at the
same rate regardless of whether they would normally be paused or slowed. At the same time, you add
radiolabeled UTP and allow transcription to occur for a short time (enough time to incorporate 20-30
nucleotides), before lysing the cells.
3.
You then isolate RNA and shear it by sonication into approx 100bp fragments and hybridise
it to a slot blot using the probes diagrammed in Fig A. You see the following result (This is a slot blot,
you have bloted probes A through G onto a nitrocellulose membrane in the shape of a slot to make things
look pretty, this is analagous to a Northern):
A B C
D E F
G
AAAA B C D E F G (i.e. the strongest band by far is A and then C is pretty strong)
How do you interpret this result? [5 points]
There is a pause site at C because after the salt increase all RNAPol transcribes and the intensity of label
is therefore proportional to the frequency of RNAPol occupancy at that point in the gene. More
occupancy means that RNAPol spends more time there. A is very strongly labeled because most RNAPol
is engaged in abortive initiation at the start site. E, F and G could be labeled more weakley because
RNAPol is aborting at D at some frequency or because the rate of elongation increases after D. Key
point: Intensity = frequency of RNAPol at a given site. Frequency is a function of # RNAPol molecules
traversing region and time taken to traverse region.
7. You run some algorithms to check for RNA secondary structure in the nascent transcript that would be
produced around the pause site. The algorithms predict a stem loop structure near the pause site. You
hypothesise that this stem loop may cause pausing of RNAPolB. Design a strategy to test this hypothesis
in vivo. [5 points]
Run the same kind of transcriptional run off assay described above on a series of mutants in which you
first mutate the putative helix to abrogate base pairing and then make compensatory mutations to restore
base pairing. Key point: Must make compensatory mutations to restore functionality of a HELIX.
Otherwise could be sequence specific or due to some other structure you hadn’t thought of.
8. From your results, the structure looks bona fide. You grow your bacteria on two types of media, A and
B. You notice that, when grown on medium B, a short RNA product can be isolated from Bac that ends
right at the pause site. No full length RNA product is detectable under theses conditions.
This short RNA is not detectable when cells are grown on medium A. You decide that RNAPolB
is terminating at this site when grown on medium B. You surmise that an anti-termination factor is
preventing this termination when grown in condition A. Devise a genetic strategy to clone this antitermination factor. [5 points]
Take the 5’ of gene X up to the position of the stem loop and place it on a vector in frame with a
selectable marker. Mutagenise the cells and grow them on medium A. Replica plate to medium A +
selection. Those cells that can’t grow are probably deficient in the anti-termination factor. Need to
control well for mutations in the selectable marker.
9. You clone the anti-terminator and name it Ant. You are curious as to its function. You determine that
this is a classII pause site. Devise an in vitro experiment to distinguish between these two hypotheses:
A)
Ant pushes the RNAPolB forward to recapture the 3’ of the RNA chain at the active site.
B)
Ant triggers an endonucleolytic cleavage to remove the RNA 3’ of the active site. [5 points]
By sequential addition and removal of single NTPs or pairs of NTPs, walk RNAPolB to a position just
prior to the pause site. Then add radiolabelled NTPs such that only the nucleotides right at the 3’ end
are labeled. Allow the polymerase to pause and backtrack. Wash out the radiolabel. Now add Ant and
cold NTPs. Where is the radiolabel? Run a gel. If it is in a short fragment, B is true. If it is in the full
length chain, A is true. Never rely on drawing inferences from a negative result.
End Problem 1
Problem 2: “Sugar, Sugar (oh, honey, honey)”…..
Your lab studies the mammalian transcriptional regulator SugR, which was identified based on homology
to other transcription factors. SugR is expressed in many tissues, though your lab is studying it in the
pancreas and the liver, where it is thought to be involved in the response to sugar intake (in order to
control insulin output and glycogen storage). Previous research indicated that stimulating pancreatic cells
with membrane-permeable cAMP causes SugR to activate the expression of the genes HonE and JelE .
Microarray analysis has indicated that gene expression of HonE and JelE is altered in a sugR mutant.
In addition, it is known that SugR is can be phosphorylated by the kinase CanD at a single residue K123,
both in vivo and vitro. In order to study the effects of SugR phosphorylation on the expression of HonE
and JelE, you decide to create a SugR K123R mutant, where the site of phosphorylation has been changed to
an arginine.
2.1 Why is it a reasonable choice to create the K123R mutation, rather than the standard alanine
substitution? What other type of mutation might you make to probe the function of the phosphorylation
site on SugR? (3 points)
My bad – I screwed this question up, so nobody was required to respond. What I had been trying to
address was the issue of making mutations that can’t be modified, but still maintain the charge of the
original residue so there is a better chance that the protein will fold correctly. Other common mutations
to make are alanine substitutions (can’t be modified) and glutamate/aspartate substitutions (to mimic the
negative charge of a constitutively phosphorylated site).
Though several domains have been identified in SugR, little is known about how SugR works to control
transcription of its target genes.
2.2a Though the microarray analysis clearly indicates that HonE and JelE are activated by SugR in , it is
not known if this is a direct or indirect effect. What technique would you use to determine if SugR is
bound to the promoters of HonE and JelE in vivo? (1 point)
ChIP – Chromatin immunoprecipitation
2.2b Please write out a flow chart of the major steps in the experiment you named (10 points)
-Collect cells (in this case, they may have been pretreated with cAMP, but this isn’t a general step)
-Crosslink proteins to everything (protein, DNA, etc) using formaldehyde.
-Lyse cells
-Shear DNA (usually get to ~500bp fragments using sonication)
-IP with antibody against SugR – will pull down SugR and anything that’s bound to it (Ab usually
conjugated to beads so they can be sedimented easily)
-Reverse the crosslinks (heat)
-Remove protein (proteinase K treatment)
-PCR w/ primers for the promoters of HonE or JelE
-Run PCR products on a gel to look for your desired products
** If your protein of interest is bound at the promoters you check, you should see an enrichment of the
promoters in comparison to the input (DNA taken after the shearing step, before IP; PCR with the same
sets of primers).
2.2c Describe one important control you should run for this experiment. What does it control for? (4
points)
Many, many controls to run… I just wanted you guys to pick one and describe it. Some biggies to think
about:
-Control for PCR primers – Run the PCR on the input DNA (kind of doing this anyway to check for
enrichment)
-Control for IP – run “IP” without adding Ab; want to ensure that any DNA amplified was pulled down
by your protein, and isn’t the result of non-specific interactions with the beads, or something random in
your buffers, etc.
-Control for IP – check specificity of antibody by using primers against positive and negative control
promoters (promoters of genes that you know are or aren’t controlled by SugR). Hopefully you won’t get
signal from your ‘negative control’ primers, b/c signal would indicate that your Ab is pulling down DNA
(or other proteins) non-specifically.
Other caveats to think about: If comparing the ChIP pulldowns with 2 Ab (like our -SugR and -SugRPO4-), need to think about relative affinities of the antibodies; and do the Ab cross-react? Also, there’s
always the possibility that your Ab recognizes the DNA binding domain, so if SugR is already bound to
the DNA, Ab won’t be able to pull down your SugR.
2.2d How would you determine if the SugR bound to the promoters is phosphorylated? (1 point)
Run IP step with an antiobody specific for the phosphorylated form of SugR (would need to run tests to
ensure that it only recognizes the phosphorylated form, and not the unphosphorylated form).
Pancreas & Liver
cAMP
Ab =
SugR
input
+
No Ab
+
+Ab
-
+
HonE
JelE
ActA (control)
HonE
Ab =
SugR-PO4-
JelE
ActA (control)
2.2e Your results are shown above. From these data, what can you conclude about SugR bound to the
target promoters before and after cAMP treatment? (2 points)
I had designed the assays such that the -SugR only recognizes unphosphorylated forms, and -SugRPO4- only recognizes the phosphorylated form. Going with this, you would conclude that in both tissues,
more SugR is bound to the promoters of both HonE and JelE after cAMP treatment. The bands are equal
sizes, so you could speculate that the binding of SugR is in no way affected by the phosphorylation state
of SugR, but this also assumes that both antibodies bind with the same affinity.
2.2f A post-doc in lab has found that the JelE gene is not expressed in liver cells when SugR is
phosphorylated. You had previously hypothesized that SugR binding at JelE was dependant on its
phosphorylation state. Instead of the results above, what results might have you been expecting? Please
redraw the results. (2 points)
Because you know that cAMP treatment causes SugR to activate the expression of JelE and HonE (see
intro paragraph), the fact that JelE is not expressed in liver cells when SugR is phosphorylated may have
led to the hypothesis that phosphorylated SugR can’t bind to the JelE promoter. Under this hypothesis,
you’d expect your ChIP data to look the same as above, except there wouldn’t be any bands for JelE
using the -SugR-PO4- antibody.
2.3. The same trusty post-doc has identified a liver-specific repressor that binds at the promoter of JelE
when SugR is also phosphorylated. Experimental data shows that the liver-specific repressor binds to a
piece of DNA located at the JelE promoter (and not at the HonE promoter). What technique would you
use to determine the exact location on the DNA where the repressor binds? Describe generally how this
technique works (you don’t have to list steps of a protocol). (5 points)
DNA footprinting; this technique is useful when looking for binding sites in a known region of DNA.
You’d isolate JelE promoter DNA, and end-label the 5’ end. Allow purified liver-specific repressor to
bind to your promoter DNA. Anywhere the protein binds, the DNA fragment will be protected against
DNaseI digestion. On a naked piece of DNA, a light digestion (such that each DNA fragment will only be
cut once), would give you a ladder of DNA fragments, spaced one bp apart on an unbound DNA
fragment. Wherever the liver-specific repressor binds to the promoter of JelE, those bands will be missing
from the ladder b/c DNaseI wouldn’t be able to cleave the DNA at the protected region.
Note – this assumes that the liver-specific repressor can bind the JelE promoter without SugR.
You now undertake a functional domain mapping experiment to determine which functional domains of
SugR are needed to differentially regulate HonE and JelE.
Three functional domains of SugR have been identified previously. They are labeled A, B and C. You
make knockout of these domains in all combinations, and perform microarrays to determine if the target
genes HonE and JelE are still being appropriately expressed in the liver. In addition, you have returned
from a quick trip to the Shokat lab with a small molecule inhibitor of CanD, which you also throw into
the mix.
To prep the cells, you treat them with cAMP to ensure SugR is bound to the target promoters. In certain
cases, the cells have been pretreated with CanD inhibitor (b/c you don’t know the order of action). The
results of your microarrays are presented below. All data was converted such that the ratio of signal is:
mutant SugR  CanD inhibitor
wt SugR, no CanD inhibitor
(
)
for each particular mutant.
(note: assume CanD is working, unless inhibited).
A “-“ indicates that expression was lower in the mutant than in the wt.
A “+” indicates that expression was higher in the mutant than in the wt.
A “0” indicates that expression was equal in the mutant and wt strains
Each mutant is indicated by the domains present. They are listed in the order (A, B, C). Any domain that
was deleted in the mutant is indicated by a 
.
JelE HonE
Inhibitor?
SugR mutants
AC
C A

C

-
-
-
-
-
-
-
-
-
0
0
0
0
0
0
0
0
0
0
0
0
ABC
AB
N
0
0
-
-
Y
0
0
-
N
0
+
Y
+
+
2.4a Please fill out the chart below, indicating which domains are needed for proper regulation of each
target gene. Place a check mark in the boxes to indicate which domains are required. Explain how the
data led to your conclusions. (7 points)
A
B
HonE
X
X
JelE
X
X
C
X
HonE: Domains A and B are both needed to get proper regulation of HonE. Without A, B, or both,
expression of HonE is decreased in comparison to wt. Domain C isn’t needed, b/c altering it doesn’t
change expression levels, and the regulation of HonE isn’t dependent on SugR’s phosphorylation state
(b/c using the CanD inhibitor doesn’t alter expression data).
JelE: Domain C is needed for proper regulation of JelE, as evidenced by the increase in expression when
only domain C is knocked out.
Domains A and B are also needed for proper regulation, b/c when you look at the A- - and the -Bmutants, you don’t get the same increase in expression as seen in the C knockout (AB-). Keep in mind
that JelE is normally not expressed in liver cells when SugR is phosphorylated, so you can’t see any lower
expression level… hence knocking things out that would be needed for expression will still give you an
“unchanged” ratio on the microarrays.
2.4b For each domain, state whether you think it’s involved in activation, repression, both, or neither.
Explain why. (6 points).
Domains A and B: activation domains. On Hone, you get lower expression when A, B or both domains
are mutated.
Domain C: repression (for JelE only): when C is gone, get increased expression. It also seems to be
involved in sensing the phosphorylation state b/c when CanD is inhibitied (row 4 of chart), wt strains
show increased expression over wt strains with the inhibitor, indicating that repression by domain C may
only work when SugR is phosphorylated.
2.4b. Based on the results, you conclude domain C is important for the alternative regulation between
HonE and JelE. One hypothesis is that this domain is an interface between JelE and the liver-specific
repressor. A biochemical and a genetic technique you could use to verify this is a contact. (2 points)
Lots of answers… for genetic, the best answer was yeast 2-hybrid. For biochemical, the best answers
were label-affinity transfer or gel shifts.
2.4c. Describe one of the techniques listed above in detail. (7 points)
Most of you got these explanations just fine, but I did want to point out a few details…
For yeast two hybrids – you’re linking the Gal4 activator domain and binding domain to domain C and
the liver-specific repressor. You’d need to do this with an isolated domain C of SugR, that can fold
correctly (assuming this is possible). If you did it with intact SugR, you couldn’t ensure that the point of
contact between the liver specific repressor and SugR was domain C. Obviously, lots of false-negatives
and positives for this technique.
For the gel shifts – The best way to use gel shifts in this context is again using only domain C and purified
liver-specific repressor. The idea is that you could run them out on a nondenaturing PA gel, and if they
associate you’d see a shift in the band. You can detect your bands using antibodies specific to each
protein (or if you’re using entirely purified proteins, just do a silver stain or Coomassie). If you use the
entire SugR protein, you again can’t be sure that the liver-specific repressor is only binding at domain C.
If you use the promoter DNA in the gel (looking for a shift when one of the proteins binds and a super
shift when both bind), you can’t be sure that the proteins aren’t binding solely to the promoter DNA; You
could get a supershift without the proteins even contacting one another.
End Problem 2
Problem 3:Rachel’s Problem (Yup, you guessed it, it’s Chromatin)
a) You’ve isolated a novel 20-protein complex from Saccharomyces cerevisiae. You decide to call it
Imaginary Protein Complex (IPC). You purify all the components, clone and sequence them, and find a
protein, Ipc1, with homology to known histone acetyltransferases.
i) 5 pts How would you test in vitro whether this protein has HAT activity? Outline your experiment and
diagram the results.
Incubate with labeled Acetyl-CoA and histones, look for transfer of label to histones by Western (look for
overlap of radioactivity and band when probed with anti-histone antibody).
ii) 5 pts What data could you obtain that would imply that this protein has HAT activity in vivo?
Knock out gene and look for general decrease in histone acetylation by western using anti-acetylated
histone antibodies (or mass spec).
b) 5 pts Among the other subunits of this complex are Ipc2, 3, and 4, proteins with homology to members
of the SWI/SNF complex, a well-known ATP-dependent chromatin remodeling complex. How would
you test whether your complex has chromatin remodeling activity? Outline your experiment and diagram
the results.
Micrococcal nuclease digestion – change in digestion pattern (in vitro – incubate purified Ipc2, 3, 4,
ATP, Mg, with a nucleosome array and look for shift in nucleosome positions).
c) 7 pts You imagine that IPC may remodel chromatin in one of three ways: by knocking nucleosomes
off the DNA so that new nucleosomes may be loaded in different positions, by sliding nucleosomes to a
different position on the same piece of DNA, or by transferring the nucleosomes to another piece of
DNA. How would you differentiate (in vitro) between these three possibilities? Hints – you don’t have
to design one experiment that will differentiate between the three, you can have a series of experiments.
Assume that you can construct any sort of DNA-nucleosome complex that your heart desires, i.e., with
any sort of sequence and nucleosome positioning.
The main difference between model 2 and models 1 and 3 is whether or not the nucleosome comes off the
DNA or not. Thus you could take labeled DNA, assemble nucleosomes on it, add IPC and excess of
unlabeled DNA and ask whether the nucleosomes come off the original piece of DNA or not. To further
get at the question of whether IPC slides nucleosomes along the piece of DNA you could construct a
nucleosome array that puts barriers to sliding on either side of the nucleosome (such as a Holliday
junction or a large tethered protein) and ask if IPC can still remodel.
The difference between models 1 and 2 is what happens to the nucleosome that gets knocked off – does it
get loaded onto another piece of DNA or not? To answer this, make a nucleosome array with labeled
histones. Add IPC and naked DNA (can label DNA with a different label) and ask whether the labeled
histones transfer to the new DNA or are just released.
d) 5 pts Just as you are about to submit your exciting results, an article comes out from a competing lab
in which it is shown that Ipc2 and Ipc17 are activated by phosphorylation in response to osmotic stress.
You do some more intensive BLASTing and find that Ipc17 as well has a slight homology to known
chromatin remodeling proteins. You decide to hold off on submitting your manuscript in the hope of
including some physiologically relevant results. First, you want to ask whether Ipc2 and Ipc17 play a role
in the regulation of the gene Osmotic Stress Reponse 1 (OSR1), whose transcription is greatly increased
in response to osmotic stress. What experiment or experiments would convince you that Ipc2 and Ipc17
are involved in regulating OSR1 transcription?
Make a reporter construct with the ORS1 promoter driving lacZ (for example) and ask whether there are
different levels of B-gal in WT vs. ipc2∆ or ipc17∆ cells.
e) Luckily for you, Ipc2 and Ipc17 are not essential. Therefore, you make the following strains: ipc2∆,
ipc17∆, and ipc2∆ipc17∆. You isolate nuclei from your WT and mutant strains in the presence or
absence of salt, treat with micrococcal nuclease, purify the DNA, and digest with restriction enzymes that
flank the OSR1 promoter. You run your DNA on a gel and probe with an end-labeled probe to the OSR1
promoter. You obtain the following results:
WT – salt WT + salt
ipc2∆ + salt
ipc17∆ + salt
ipc2∆ipc17∆ + salt
Naked DNA
i) 6 pts What are two possible models for the functions of Ipc2 and Ipc17 that explain these digestion
patterns?
1. Ipc17 knocks off nucleosomes and Ipc2 puts them back on.
2. Ipc17 “disorders” nucleosomes and Ipc2 reorders them.
ii) 4 pts What experiment could you perform that would distinguish between these possibilities?
In model 1, there are no histones at the ORS1 promoter in an ipc2∆. In model 2, histones are still present
but in different positions. Do a ChIP using anti-histone antibodies at the ORS1 promoter in WT vs ipc2∆.
f) 6 pts You are a huge fan of David Allis and think that maybe he would give you his autograph if only
you could decipher the histone code. You decide to start small. You hypothesize that Ipc1 may acetylate
certain residues of histones at ORS1 that would increase binding by other Ipc proteins. You decide to
perform a screen for histone residues that are involved in ORS1 transcription. Tell me about it –
remember to include details of your strain genotypes, read-out/selection for the screen, and remember that
all histone proteins are essential.
Make a strain that has whatever histone you’re testing knocked out and covered by a wild-type copy on a
plasmid (say Lys). Put in a reporter construct with the ORS1 promoter driving a Ura gene. So under
osmotic stress, these cells presumably can’t grow on FOA since the Ura reporter is being expressed. Do
sloppy PCR on another plasmid containing the histone, shuffle it into the strain and get rid of the wildtype plasmid, and look for mutants that grow on FOA under osmotic stress. Sequence them.
g) 7 pts You find that Lysine 14 of histone 3 is important for transcription of ORS1. Through a
modification of your original experiments you find that Ipc1 acetylates H3K14. You come up with three
different models for the effects of IPC on ORS1 transcription: A) Ipc1 acetylates H3K14 at the ORS1
promoter, the chromatin-remodeling Ipcs recognize acetylated H3K14 and do their chromatin-remodeling
thing at the ORS1 promoter, which allows transcription factors to bind and activate ORS1 transcription.
B) The chromatin-remodeling Ipcs act first to remodel chromatin at the ORS1 promoter, which allows
Ipc1 to bind and acetylate H3K14; transcription factors recognize acetylated H3K14 and activate
transcription from the ORS1 promoter. C) Acetylation of H3K14 and chromatin remodeling happen in
concert to promote transcription. Design an experiment (or experiments) to differentiate between these
possibilities, keeping in mind that you don’t know the specific transcription factors that activate ORS1
transcription. For the purposes of this question, assume that the order of chromatin remodeling events
described in part e) for Ipc2 and Ipc17 is independent of other events (i.e., just think about chromatin
remodeling as one step).
1) Knock out Ipc1 (or make a non-acetylatable H3K14 mutant) and ask whether the chromatin
remodeling Ipcs can still bind ORS1 promoter (do ChIP using anti-Ipc2 antibody for ORS1 promoter, and
also do nuclease digestion assays on ORS1 promoter to see if you still get remodeling or not).
2) Knock out Ipc2 and ask whether Ipc1 can still bind and whether H3K14 still gets acetylated or not.
If 1 is true, then it’s model A. If 2, then model B. If neither, then C.