Download 7.06 Problem Set #5, Spring 2005

7.06 Problem Set #5, Spring 2005
1. You are working as a researcher for a biotechnology firm, developing orange trees
that are more resistant to cold weather. There is a class of cold-response proteins
called COR proteins that you have been studying. Wheat, which grows under widely
different climates, expresses a particular COR protein (WheCOR) that is not expressed
in orange trees. The WheCor gene is nuclear-encoded, and the WheCOR protein is
targeted to the chloroplast stroma. You hypothesize that making a transgenic orange
tree that expresses WheCor might make it more resistant to cold weather.
a. First you decide to determine if the WheCOR protein can be transported into orange
tree chloroplasts. A colleague suggests that you do an in vitro post-translational uptake
assay. In this assay he suggests adding purified radiolabeled WheCOR made in a cellfree system to orange tree chloroplasts and some cytoplasmic extract. As a negative
control, he suggests repeating that same experiment, but adding cyanide prior to adding
the WheCOR protein. Then he says you can add the protease trypsin to both the
cyanide-treated and untreated chloroplasts. Finally you would run an SDS-PAGE gel of
your reaction mixture and ascertain if your protein is protected by the chloroplasts by
looking for signal or absence of signal in the resulting autoradiographic film. You
explain to your colleague that he is confused and that this experiment wouldn’t proceed
as he anticipates it would. What mistake is your colleague making?
You cannot use cyanide-treated chloroplasts as a negative control. Cyanide poisons the
oxidative phosphorylation pathway by dissipating the proton-motive force. In
mitochondria, the dissipation of the proton-motive force would cause proteins to be unable
to enter the organelle. However, this force only drives protein import into mitochondria,
and not into chloroplasts.
b. After talking to your confused colleague, you decide to design your own experiment.
Therefore you introduce the WheCor gene into orange tree plant cells, using a vector
that integrates into the nuclear DNA. You observe through fluorescence microscopy
that the WheCOR protein is not being transported into the transgenic orange tree
chloroplasts, although the endogenous orange tree chloroplast proteins are. To
determine what is wrong, you do the following experiment. You use three experimental
groups: 1) wild-type wheat, 2) mutant wheat that contains a point mutation in the
WheCOR stromal-import sequence, and 3) the transgenic orange tree expressing
WheCOR. You separate the chloroplasts from the rest of the cell components (cytosol,
other organelles and membranes) for each experimental group. You then make a
lysate from both the chloroplasts and the non-chloroplast cell components, and perform
Western blotting using an anti-WheCOR primary antibody. You see the following result:
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Wheat
Rest of
the Cell
Chloroplasts
Mutant Wheat
Rest of
the Cell
Chloroplasts
Transgenic Orange Tree
Rest of
the Cell
Chloroplasts
Give a possible explanation for this result.
These results confirm that WheCOR is not being transported into the orange tree
chloroplasts. Furthermore, these results show that WheCOR in the orange tree cytoplasm
is the same size as the WheCOR protein after it has had its stromal-import sequence
cleaved off in the wheat chloroplast stroma. A possible explanation for this is that the
orange tree cell contains a cytosolic protease that prematurely cleaves the N-terminal
WheCOR stromal-import signal prior to entering the chloroplast. This would explain why
WheCOR is not properly imported into orange tree chloroplasts.
c. To avoid any further complications with chloroplast targeting of the protein, you
decide to introduce the WheCor gene (minus the part that encodes the stromal uptake
sequence) into your orange tree cells using a vector that integrates directly into a
specific site in the chloroplast DNA via homologous recombination. This vector contains
a selectable marker that encodes BADH, an enzyme that converts toxic betaine
aldehyde (BA) into nontoxic glycine betaine. You introduce your vector containing the
WheCor gene into orange tree plant tissue and culture the plant tissue in regeneration
medium containing BA.
As an experienced researcher, you know that introducing genes into the chloroplast
genome is trickier than introducing DNA into the nuclear genome, and that multiple
rounds of selection are required during the regeneration process for a successful
transformation. Give an explanation for why this might be.
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Each plant cell contains many chloroplasts, and each chloroplast contains multiple copies
of the chloroplast genome. When integrating a new gene into the chloroplast genome, not
every chloroplast will receive the new gene, and within each chloroplast, not every genome
will receive the new gene. This is called heteroplasmy. As chloroplasts divide, the
chloroplast genomes segregate randomly. As the cells divide, their chloroplasts divide and
are segregated randomly. Such random segregation could lead to loss of the WheCOR
transgene in future progeny. Therefore multiple rounds of selection are used to give a
growth advantage to those cells that happen to receive more chloroplasts containing more
copies of the selectable marker. This will eventually lead to homoplasmy and successful
complete transformation.
d. After successfully transforming the WheCor gene into the chloroplasts of your orange
tree, you present your work to your boss. She tells you that she is very impressed that
you thought to transform the chloroplasts. She thinks that using chloroplast
transformations will help ease the fears of environmentalists who oppose genetic
modifications to the nuclear genome of flowering plants. These environmentalists are
very concerned with the spread of transgenes from transgenic plants to wild plants
through airborne pollen or through pollination by insects. Why might chloroplast
transformations be preferable to these environmentalists?
Nearly all of a flowering plant’s chloroplasts are inherited maternally (just as with
mitochondria). There would be less fear of spreading the transgene to wild plants via
pollen (the plant equivalent of sperm) if the transgene was integrated into the chloroplast
DNA, since pollen has almost no chloroplasts.
2.
Later on in 7.06, we will be discussing a type of mammalian cell called embryonic
stem cells (ES cells). ES cells are special because they can continually grow and divide
on culture plates without “differentiating” (ceasing cell division and assuming
characteristics of cells that have chosen to carry out specific functions). However, when
ES cells become too dense on a culture plate, they do start differentiating. You discover
a gene DIFF1 (Differentiation1) that, when deleted, results in the inability of mutant ES
cells to differentiate in response to density. Intrigued by this system, you study the
protein encoded by DIFF1. You speculate that it may be a transcription factor that
translocates into the nucleus of ES cells that have sensed that they are surrounded by a
high density of other cells.
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a. You develop an antibody to DIFF1. How would you use the antibody to test your
hypothesis, and what results would you expect to see?
You could do immunofluorescence experiments in which you would visualize DIFF1 in cells
growing on high density and low density plates. If DIFF1 indeed translocates into the
nucleus when some critical density of cells on the plate is achieved, all of DIFF1 antibody
staining should be distinctly nuclear in crowded cells. When you perform
immunofluorescence on low density cells, DIFF1 staining should be cytoplasmic. You can
also stain the total DNA in these cells with a dye that fluoresces in a different wavelength
than your secondary antibody to be sure that what you see is truly nuclear or cytoplasmic
staining.
b. Your experiment works and you have evidence that DIFF1 indeed translocates into
the nucleus upon crowding. How would you show that DIFF1 can also then shuttle back
out of the nucleus when growth conditions change? (Make sure to design an
experiment that ensures that the DIFF1 you see in the cytoplasm is not just newly
synthesized DIFF1.)
Visualing DIFF1 in cells grown at high density that are moved to a new low density plate
won’t work, because you will not know if the DIFF1 observed in the cytoplasm was
synthesized de novo (and the nuclear DIFF1 degraded) or if the nuclear DIFF1 was
actually shuttled out. However, you could treat your cells with a protein synthesis inhibitor
such as cyclohexamide and see if, after being transferred for an hour or two into a low
density environment, cells display staining of DIFF1 back in the cytoplasm. If you start
seeing DIFF1 in the cytoplasm, you can conclude that DIFF1 also shuttles out of the
nucleus.
Alternatively, you can use a heterokaryon assay. For this, you would need to fuse high
density DIFF1+ ES cells with DIFF1-/- ES cells. During cell fusion, treat cells with
cyclohexamide (or another reagent which inhibits protein synthesis). After an hour or two
you can then visualize DIFF1 by immunofluorescence. If you see DIFF1 present in the
DIFF1-/- nucleus of a heterokaryon, then you can conclude that DIFF1 can shuttle both
into and out of a nucleus. Please note that, in order to do this experiment, you need to have
some sort of a marker that will tell you which nuclei in the heterokaryons are DIFF1-/-,
and which nuclei in the heterokaryons are DIFF1+.
c. You successfully find that DIFF1 can be exported back out of the nucleus if crowding
is eliminated. Knowing that DIFF1 can be both imported into and exported out of the
nucleus, you examine the amino acid sequence of DIFF1 for NES and NLS sequences.
You find that DIFF1 protein contains two putative NLS’s in its sequence, and you name
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them cluster A and cluster B. You suspect that only one of those clusters is actually
necessary for nuclear transport. How would you test this hypothesis?
Use site specific mutagenesis to engineer DNA constructs that express mutant forms of the
DIFF1 protein which either lack cluster A or cluster B. Express these mutant forms of
DIFF1 in DIFF1-/- cells. Then examine by immunofluorescence whether DIFF1 without
either cluster A or cluster B still translocates into the nucleus after high density growth is
achieved. If “DIFF1w/oA” appears to translocate into the nucleus, but “DIFF1w/oB” does
not, then you know that cluster B was necessary for nuclear translocation.
d. Once you figure out which cluster is necessary for DIFF1’s ability to translocate into
the nucleus, how would you test if this cluster is also sufficient for nuclear import?
Engineer a DNA construct that will encode a fusion of a protein which is known to be
always cytoplasmic to DIFF1’s NLS sequence. Use immunofluorescence against the
resident cytoplasmic protein to examine if this protein now has the ability to translocate
into the nucleus. If it can go into the nucleus, then DIFF1’s NLS is sufficient for nuclear
import.
e. You perform a series of co-immunoprecipitation experiments at different time points
after ES cells are made to feel crowded. In your experiment, you induce crowding and
inhibit protein synthesis at the same time. You then immunoprecipitate DIFF1 at
various timepoints after crowding has been induced. You find that, one hour before
differentiation is first observed, Importin can be co-immunoprecipitated with DIFF1.
However, after increasing amounts of time after crowding, you cease to be able to coimmunoprecipitate Importin with DIFF1. How would you explain this?
Importin binds to DIFF1 and brings it into the nucleus through the nuclear pores.
Subsequently, in the nucleus, a GEF protein facilitates the conversion of Ran-GDP to RanGTP. Ran-GTP binds Importin with high affinity and displaces the cargo protein DIFF1.
Thus, as more and more DIFF1 translocates into the nucleus with increasing cell density,
you observe less and less Importin complexed with DIFF1. When all of cell’s DIFF1 is in
the nucleus, you will no longer see it interacting with Importin at all.
f. You decide to examine DIFF1’s location in cells that are overexpressing a mutant
form of Ran that cannot hydrolyze GTP (even in the presence of GAP proteins). Where
do you expect to see DIFF1 in these cells when they were grown at high density, and
why?
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All of the DIFF1 would eventually end up in the cytoplasm. This is because Ran-GTP is
normally high only in the nucleus. Ran-GTP interacts with importins and causes them to
release their cargo in the nucleus. If Ran that cannot hydrolyze GTP is expressed
throughout cells, then Ran-GTP becomes high in both the cytoplasm and the nucleus.
Therefore, importins are always bound to Ran-GTP, and never bind their cargo.
Therefore, any cargo that is supposed to be delivered to the nucleus won’t get there.
Additionally, Ran-GTP interacts with exportins and causes them to bind to their cargo in
the nucleus. This trimeric complex then shuttles into the cytoplasm. If all Ran in the cell is
Ran-GTP, then exportins will always be bound to DIFF1, and will constitutively bind and
export their cargo. A combination of the inability to import DIFF1 and the constitutive
ability to export DIFF1 will eventually lead to all DIFF1 being in the cytoplasm.
3. Many cell types within multicellular organisms are motile and can move around
within a tissue. Motile cells secrete special enzymes to accomplish this task. These
secreted enzymes allow motility by chewing up the gooey extra-cellular matrix (ECM) in
which cells are surrounded. In the hopes of identifying a new secreted enzyme that
digests ECM, you have chosen to study a gene of unknown function whose expression
is upregulated in mammalian cells that have an enhanced ability to move through an
ECM.
a. You confirm experimentally that your protein is targeted to the secretory pathway.
Next, you are interested in whether or not your protein is a transmembrane protein.
How can you determine this using microsomes in a cell-free system?
In a cell, if your protein is secreted, after being targeting to the ER membrane, it will be cotranslationally translocated into the ER lumen. From there, the protein will travel within
ER and Golgi vesicles to the plasma membrane, and will be exocytosed. If the protein is a
plasma membrane, ER membrane, Golgi membrane, or lysosome membrane protein, it
will be partially translated into the lumen of the ER, but a segment of the protein will cross
the ER membrane, and a segment of it will remain outside of the ER in the cytoplasm. As
the protein travels through the secretory system, it will eventually reach its target
membrane via vesicle-to-membrane fusion, and the cytosolic portion of the protein will
remain cytosolic.
Microsomes are made by disrupting the ER to create small ER vesicles that can act like a
normal ER in vitro. If you add microsomes to ribosomes and the mRNA encoding your
protein to a test tube, the mRNA will be co-translationally translocated into the microsome
as explained above. You then treat the microsomes with proteases, extract your protein
from the microsomes using detergent, and run your protein on an SDS-PAGE gel. If your
protein is shorter than the size it is when the same experiment is conducted in the absence
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of protease treatment, that means that your protein was cleaved by the proteases. Thus
part of your protein must have been exposed (not inside the microsomes) and susceptible to
the proteases. That would indicate that the protein was a trans-membrane protein,
because the cytosolic segment of the protein was not protected from the proteases by the
microsomes. If your protein is the same size regardless of whether you treated with
protease or not, that would indicate that your protein was fully inside the microsome, and
thus not exposed at all to the protease. This would indicate that your protein was not a
transmembrane protein, but rather destined to be either a secreted protein, an ER lumen
protein, a Golgi lumen protein, or a lysosome lumen protein.
b. You find out that your protein is a transmembrane protein. To further study the
mechanism of translocation of your protein into the ER, you decide to use microsomes
containing the SRP receptor and the Sec61 complex (the translocon). You compare
the rates of translocation of your protein from two different experiments. In the first
experiment, you add the mRNA encoding your protein to a cytosolic extract containing
ribosomes and you add microsomes to this test tube. In the second experiment, you
use in vitro translation to generate your protein, and then you add your protein to a tube
containing microsomes and cytosolic extract. How do the rates compare between the
two experiments, and why?
Your protein would only be able to be translocated when it is being translated at the same
time. Therefore, the first experiment would result in translocation, but the second
experiment would not. Proteins in mammalian cells that are destined for the secretory
pathway are always co-translationally translocated into the ER. During co-translational
translocation, the energy needed to “push” proteins through the translocon is actually
provided by the elongation process on the ribosome.
c. A colleague of yours is working on protein translocation in yeast. She tells you that,
when one does the same type of comparison as was done in part b) using yeast
proteins and yeast microsomes, a different result from what was obtained by you in part
b) can be obtained. How and why is that possible?
In yeast, some proteins destined for the secretory pathway can be post-translationally
translocated. Therefore, your colleague has conducted translocation experiments in yeast
that do work when the protein is translated first, before microsomes are added. Since the
ribosome is not around to supply the energy for translocation once the protein is fully
translated, chaperone proteins (such as BiP) are required to “pull” the protein through the
translocon from the inside of the ER membrane (or liposome). Additionally, since the
ribosome is not required for post-translational translocation into the ER, neither SRP nor
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the SRP receptor are required for translocation. Note that this post-translational
translocation process does require another Sec complex called Sec63.
d. Since your protein is a transmembrane protein, you want to see what class of
transmembrane protein it is: Type I, II or III. Come up with an experiment using
microsomes that will narrow down your options. Available to you are two monoclonal
antibodies, one of which is specific to the N terminus of your protein, and the other of
which is specific to the C terminus of your protein.
Type I, II, and III proteins have one membrane-spanning domain. Type II proteins have
their N-terminal domain in the cytoplasm and their C-terminus in the ER lumen. Type I
and III proteins have the opposite topology. To distinguish between these two groups, you
can add microsomes to a solution of ribosomes translating the mRNA of your protein (as
discussed in part a). After translocation has occurred, you can treat one tube of this
solution with an antibody against the N-terminus of your protein and one with an antibody
against the C-terminus of your protein (in the absence of detergent, so the microsomes
remain intact). You then wash away free antibody, and detect whether or not antibody
remains bound to your protein. If the anti-N-terminal antibody can bind to the protein
after translocation into the microsomes, then the N-terminus of your protein is cytosolic
and your protein must be a Type II protein. If the anti-C-terminal antibody can bind to
your protein after translocation into the microsomes, then the C-terminus of your protein
is cytosolic, and your protein is either a Type I or III protein.
e. From the experiment in part d), you narrow down your protein to either being a Type
I or a Type III protein. How might examining the amino acid sequence of your protein
by eye (or using a computer) help you distinguish between these two possibilities?
The typical signal sequence that targets a protein initially to the ER, and thus to the
secretory pathway, is a 16-30 AA long sequence that contains one or more positively
charged residues adjacent to 6-12 hydrophobic residues. Type I proteins contain this signal
sequence at their N-termini. In contrast, Type III proteins lack these cleavable N-terminal
ER sequences, and instead contain a single internal hydrophobic sequence that functions
both as the ER signal sequence and a membrane-anchor sequence. Thus you could use a
computer program that searches for N-terminal signal sequences, and conclude that your
protein is most likely Type I if it contains an N-terminal signal sequence.
4. You are studying the secretory pathway in cultured mammalian cells.
You are
interested in the retention of ER proteins in the ER despite the large number of vesicles
that are constantly budding off from the ER and traveling to the Golgi. ER resident
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proteins that are present in high concentrations in the ER are often passively
incorporated into the COPII vesicles that shuttle from the ER to the cis-Golgi. To
prevent depletion of these important ER proteins, many of them contain a C-terminal
KDEL sequence that allows them to be retrieved back to the ER by binding to KDEL
receptors present in COPI vesicles that travel from the Golgi back to the ER. KDEL
receptors are also present in COPII vesicles heading from the ER to the cis-Golgi,
because these receptors need to be returned to the Golgi to do their job.
a. Importantly, proteins bearing the KDEL sequence preferentially bind to KDEL
receptors found in COPI vesicles (and don’t bind to KDEL receptors found in COPII
vesicles). Design an experiment to show that the pH differences between COPI and
COPII vesicles provide the mechanism for this preferential binding. (Keep in mind that
KDEL receptors are transmembrane proteins that must be maintained in a membrane to
retain their proper structure and function.)
One approach is to perform binding assays with purified KDEL receptors embedded in
liposomes. These receptors would have to all be incorporated into the liposomes in the
reverse orientation of how they are in vesicles in the cell so that they can access the KDEL
ligands in the solution. By adding a radioactive peptide or protein containing the KDEL
sequence at increasing concentrations at the pH of the ER versus the pH of the cis-Golgi,
one can show that the KDEL receptor has a lower Kd for its ligand at the cis-Golgi pH
(which would be the pH of the COPI vesicles). The affect of pH on the Kd provides a
mechanistic explanation for how the KDEL receptor can be present in both the COPI and
COPII vesicles, but preferentially bind to its ligand in COPI vesicles.
b. You have isolated a mammalian cell line that is homozygous for a temperature
sensitive mutation in the gene that encodes COPI. COPI proteins are required for
retrograde trafficking of vesicles from the cis-Golgi to the ER. In order to study how this
affects transport of proteins in the secretory pathway, you grow cells expressing the
viral protein VSVG at the permissive temperature, and then shift cells to the nonpermissive temperature. After several hours at the non-permissive temperature, you
feed the cells a short pulse of radioactive amino acids to label any newly synthesized
proteins. After 30 minutes, you extract the viral protein VSVG from the cells, which is a
cell surface protein when it is properly made. You treat the extracted protein with the
enzyme endoglycosidase D, which cleaves oligosaccharide chains from proteins that
have been processed in the cis-Golgi, but does not cleave proteins that remain in the
ER. As a consequence of this cleavage, VSVG from the cis-Golgi migrates at a faster
rate during SDS-PAGE than VSVG from the ER.
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Draw into the lane labeled “ts cells” what you would expect to see when you run out
VSVG from mutant cells on a gel. Explain your answer.
wt
cells
Resistant
Sensitive
ts
cells
?
You would expect to see the VSVG band from ts cells to run at the resistant length only.
This is because loss of functional COPI proteins will disrupt retrograde trafficking of key
ER proteins (such as v-SNARES) that have moved on to the Golgi. Many of these proteins
are necessary for proper anterograde trafficking from the ER to the cis-Golgi, but in the
COPI-ts mutant, these proteins will not be returned to the ER for re-use. Over time,
depletion of such proteins will thus prevent anterograde trafficking, leading to retention of
newly synthesized proteins such as VSVG in the ER.
c. The maturation of the Golgi (from cis to medial to trans) is thought to occur by
cisternal progression. This model requires that resident Golgi proteins be returned to
less mature (earlier) Golgi stacks from more mature (later) Golgi stacks by COPI
vesicles. In this model, retrograde vesicle transport would actually cause the maturation
of the Golgi. An alternative model of Golgi maturation is that maturation occurs
because the vesicles seen traveling between stacks are carrying secretory pathway
proteins from earlier stacks to later stacks. In this model, stacks remain either cis or
medial or trans, and secretory pathway proteins are brought from stack to stack.
Outline an experiment to distinguish between these two models (vesicular transport and
cisternal progression). Your experiment should involve performing immuno-EM on cells
engineered to express VSVG. You have available to you gold-labeled antibodies
against COPI, COPII, VSVG, and the Golgi resident protein mannosidase II. (Note: you
may use more than one antibody at a time in immuno-EM by labeling each different type
of antibody with a different diameter of gold particle.) Explain which results you would
expect to see from the experiment if each of the two models was true.
Perform immuno-gold labeling of the ER, Golgi, and vesicles from sectioned cells, and
visualize using electron microscopy. When you stain with gold-labeled antibodies against
these four proteins:
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If cisternal progression is correct, you should not see COPII-stained vesicles containing
mannosidase II and/or VSVG traveling between Golgi stacks. This is because, in the
cisternal progression model, anterograde vesicle transport does not occur from earlier to
later Golgi stacks.
If the vesicular transport model is correct, then you should see COPII-stained vesicles
containing VSVG and/or mannosidase II traveling between Golgi stacks. This is because,
in the vesicular transport model, anterograde vesicles would be carrying secretory pathway
proteins from earlier to later Golgi stacks.
Please note that actual data obtained from cells favors the cisternal progression model.
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