Oct 112010

ResearchBlogging.orgClostridium difficile is an intestinal pathogen that causes diarrhea in hospitals and other healthcare settings (including nursing homes). Present as a commensal bacterium in a significant fraction of the population, C. difficile is usually rather harmless, its numbers suppressed by competition with the intestinal flora. When its competitors are decimated by antibiotics, however, C. difficile flourishes, releasing toxins that cause inflammation and diarrhea, which can be dangerous because the individuals suffering these effects are often already ill. There has been conflicting information, however, as to which of C. difficile‘s toxins are necessary to cause disease. A paper in the recent Nature (1) aims to resolve the question.

The two best-characterized C. difficile toxins (TcdA and TcdB) have the same general arrangement and function (and ~45% identical AA sequence). An N-terminal glucosylating domain attacks the cytoskeleton of host cells by inactivating Rho GTPases, a C-terminal domain mediates binding and uptake by the host cells, and a protease domain in the middle releases the glucosylating domain to do its work. Since these proteins appear to serve redundant functions, one might expect that both would support virulence. However, preceding work in the field has variously identified TcdA or TcdB as a key virulence factor (2,3). Differences in methodology and materials have contributed to the confusion, in part because different kinds of cells seem to be more or less susceptible to particular toxins, and different strains of C. difficile might have different behaviors.

Kuehne et al. aim to relieve some of the confusion by removing a subset of these confounding factors. In a single strain of C. difficile they inactivated the genes for either TcdA, TcdB, or both by inserting introns into them. An intron would be no problem for a eukaryote, but bacteria can’t handle them, so this has the effect of eliminating the expression of the gene. They then tested the toxin mixtures shed by the bacteria against cultured human and monkey cells. As expected, A-B- bacteria (with both toxins knocked out) showed no toxicity towards the cells, but A-B+ and B+A- variants were toxic towards both kinds of cells to roughly the same degree. This suggests that both toxins are sufficient for virulence.

This implication was largely backed up by a subsequent experiment in hamsters. The animals were dosed with an antibiotic and then infected with C. difficile spores of a single strain. Colonization occurred (in every case but one) within three days. The hamsters infected with A-B- C. difficile remained asymptomatic until the end of the experiment, but the recipients of the other strains all perished within a week. The A+B- group survived somewhat longer, but not dramatically so; again, this supports the interpretation that both proteins are sufficient for virulence.

This contrasts with an earlier study published in Nature (2) where it was shown that deletion of the B toxin protected hamsters from C. difficile-associated disease, using very similar protocols. Kuehne et al. attribute the differences in their results to the hamsters or genetic variation in the C. difficile strains used. While the virulence of the B- strain in this experiment was slightly attenuated, all colonized hamsters still died in relatively short order, and in human beings the situation might well be reversed, since cultured human cells are more vulnerable to toxin A.

The results of Kuehne et al. largely agree with earlier experiments (3) and with what one would naturally expect of two very similar toxins being released by the same organism. While susceptibility to a particular toxin may vary with characteristics of the host species or cell type, it seems likely that both toxins are capable of supporting virulence. While it is to be hoped that additional research will clarify the reasons for the discrepancy between these two experiments, efforts to treat C. difficile-associated disease by attacking the toxins should proceed with the assumption that both must inactivated. Thanks to their functional and sequence similarity this will hopefully not be too much of a complication.

1. Kuehne, S., Cartman, S., Heap, J., Kelly, M., Cockayne, A., & Minton, N. (2010). The role of toxin A and toxin B in Clostridium difficile infection Nature, 467 (7316), 711-713 DOI: 10.1038/nature09397

2. Lyras, D., O’Connor, J., Howarth, P., Sambol, S., Carter, G., Phumoonna, T., Poon, R., Adams, V., Vedantam, G., Johnson, S., Gerding, D., & Rood, J. (2009). Toxin B is essential for virulence of Clostridium difficile Nature, 458 (7242), 1176-1179 PMCID: PMC2679968 OPEN ACCESS

3. Voth, D., & Ballard, J. (2005). Clostridium difficile Toxins: Mechanism of Action and Role in Disease Clinical Microbiology Reviews, 18 (2), 247-263 PMCID: PMC1082799 OPEN ACCESS

Aug 262010
ResearchBlogging.orgIf you’re going to study the role an enzyme plays in a biological pathway, it’s often useful to “kill” it with a mutation. For example, the proline cis-trans isomerase cyclophilin A (CypA) needs a particular arginine residue for its chemistry, so mutations that remove or alter that functional group, like R55K and R55A, should destroy the protein’s function and have effects on the related pathways that help illustrate its role. The hydrophobic pocket it uses to bind substrates is made by residues like H126, F113, and W121. Growing or shrinking those residues should alter the shape of the pocket and change binding or activity, leaving the enzyme “dead”.

Using model reactions and various binding assays, researchers have previously examined a number of these mutants (4,7) and found that they diminish isomerase activity and alter inhibition. However, a detailed study of the effects of the mutations on CypA’s catalytic cycle has not been performed. Former Kern lab members Daryl Bosco (now a professor at UMass Medical) and Elan Eisenmesser (now at UCHSC) examined these mutants in greater detail to see how they really behaved. I also contributed some data at the last minute, when the third reviewer requested we study an additional mutant, prompting a scene that I promise was not too much like that Downfall parody. In every case we found that these enzymes, although significantly impaired, weren’t as dead as they had seemed.

You had me at “dunno”

CAN bound to CypA, from PDB structure 1AK4 (5).
CypA residues are labeled in black, CAN residues in red.

One key aspect of this work is that it involves a physiological substrate of CypA, namely the N-terminal domain of the HIV-1 capsid protein (CAN). Mature HIV-1 virions contain CypA that is bound to proline 90 of CAN. The absence of CypA dramatically reduces their ability to infect their target cells, which we know from experiments with mutant CA proteins as well as ones involving the CypA inhibitor cyclosporin (3). What we don’t know about the system is exactly what CypA does for HIV-1. The crystal structure (right) of CAN in complex with CypA appears to only capture the trans isomer configuration (5), but for reasons I have discussed previously on this blog, that’s not particularly informative. We know, largely from Daryl and Elan’s previous research on the system (2), that when CAN is floating free in solution CypA will catalyze isomerization, but in the context of a fully assembled capsid that situation could conceivably change.

This leaves us with three possibilities for CypA’s function in the capsid. Catalysis of cis-trans isomerization of the proline bond could be important. Or, maybe all capsid needs is for CypA to bind at P90, and catalysis is irrelevant. And perhaps neither of these functions matters and CypA just needs to be hanging around for some other reason. To address these possibilities, Saphire et al. performed an elegant series of experiments where they sneaked an engineered CypA protein into another part of the capsid by fusing it to a protein called Vpr. When they replaced the normal CypA sequence with a mutant (H126A) that was supposed to abrogate both binding and catalysis, HIV-1 could still infect CD4+ cells (6). But, how sure can we be that H126A, or any other mutant, is actually “dead”?

You can’t measure what you can’t see

The problem with proline isomerization, from a biochemist’s standpoint, is that it’s a difficult reaction to detect. While switching between isomerization states may have structurally significant effects, there’s no direct spectroscopic signal to tell you whether a proline bond is in the cis or trans conformation. Even if there was, most proteins have many prolines and so the signal of the bond you care about might be difficult to separate from the bonds you don’t.

You can get around this difficulty using a coupled reaction with a model substrate. CypA catalyzes the isomerization of tetrapeptides of the form AXPF pretty efficiently. As it turns out, sequences like this are also good substrates for the protease chymotrypsin, but there’s a catch. Chymotrypsin only cleaves substrates where the proline bond is in the trans configuration. So, what you can do is take a substrate like succinyl-Ala-Ala-Pro-Phe-p-nitroanilide, add a tiny amount of cyclophilin, and then dump in a huge amount of chymotrypsin. With enough chymotrypsin, the peptide that’s already trans will be cleaved before the solution stabilizes, causing a color change (due to the pNA) that can be measured with a conventional spectrophotometer. Then you can monitor the conversion of the remaining substrate from cis to trans, because there’s so much chymotrypsin that cleavage after isomerization is essentially instantaneous.

This works reasonably well, but it has some limitations. You’re stuck with a model peptide that may not behave very much like your particular protein substrate. You’re only following the cis-to-trans reaction, and even that comes with limited detail. Also, performing the experiment takes some careful work, because if you add too much of your CypA the reaction will end before the solution turbulence settles, and if you add too little, the intrinsic cis-trans isomerization will interfere with your catalytic measurement.

Although proline isomerization is a difficult reaction to follow by spectrophotometry, it’s actually quite convenient to assay by NMR. Because CypA catalyzes the reaction in both directions, it’s impossible to exhaust the substrate. The kinetics can therefore be measured at equilibrium using NOESY and ZZ-exchange experiments (2). Of course the experiment is limited by our ability to express isotopically-labeled substrate proteins, but provided we can do that and visualize the active site in our spectra, then we can observe catalysis of the native substrate. When you perform this experiment on these various “dead” forms of CypA using CAN as a substrate, it becomes evident they’re still active after all.

Night of the living “dead” enzymes

Panels B-G of Figure 3 in this paper directly show that every single one of the CypA mutants catalyzes CAN isomerization in solution (1). These spectra show peaks representing the chemical shifts of the nitrogen and hydrogen atoms of CAN‘s backbone amide groups in the presence of a small amount of CypA, so we are not looking at P90 directly. Fortunately, the chemical shift of the G89 amide is dependent on the isomerization state of P90. If the G89-P90 bond is in trans, G89 shows up as the large peak at lower right in these panels, but if the bond is in cis you get the small peak at upper left.

If you don’t wait very long between determining the 15N chemical shift (y-axis) and the 1H chemical shift (x-axis), you get something that looks like panel A. If, however, you pause between determining the 15N shift and the 1H shift, you get cross-peaks representing the portion of CAN proteins that started the experiment in trans and ended it in cis, or vice-versa. The presence of these cross-peaks in the CAN/CypA samples, and their absence in the CAN-only sample (panel A), proves that catalysis is occuring. I’ve blown up the figure for H126A on the right to make things a little clearer. In this case the cross-peaks were pretty weak, but still in evidence.

There was more variability when it came to affinity, the strength with which CypA binds the CAN substrate. I’ve shown a complete titration for H126A on the left. As you can see, progressive addition of H126A causes the free CAN peaks to disappear while the new bound CAN peak grows in. This behavior is characteristic of slow chemical exchange on the NMR timescale, and indicates a high-affinity binding interaction. WT CypA binds CAN with a KD of 13 µM, and H126A probably has similar affinity. Note also that the bound state has a single peak for each residue, while the free state of CAN has separate cis and trans peaks. This indicates that the cis and trans isomers are interconverting rapidly on the enzyme, and constitutes additional evidence that H126A CypA is catalytically active.

This pattern was not repeated for all the mutants, however. H126A and W121Y had affinity similar to WT, while R55A, R55K, and F113W had significantly higher KD (lower affinity). You can see this clearly from the titrations in Figure 5. For each of these mutants, adding CypA to CAN caused the CAN peaks to move around in the spectrum, rather than disappearing and reappearing (R55K had a mixture of behaviors because the NMR timescale also depends on chemical shift). This peak shifting is characteristic of fast chemical exchange on the NMR timescale and indicates relatively low affinity. This wasn’t the only change for those mutants.

Shifts in the rate-limiting step

An enzyme that doesn’t have very high affinity for its substrate isn’t necessarily in trouble. The NMR titrations of the R55A and R55K mutants indicate that their KDs are near 1 mM (Table 1). This is comparable to the affinity the WT protein has for the AAPF peptide, which gets catalyzed pretty efficiently. What does seem strange about this result is that the ZZ-exchange spectra are very similar.

The presence of single peaks for residues of CAN bound to WT suggests that the isomerization step is fast. Using relaxation-dispersion techniques, Daryl established that the net process rate (kct+ktc) for CAN on WT CypA was about 2200 /s. From the ZZ-exchange spectra we know that the total catalytic cycle goes a great deal slower (closer to 75 /s), from which we can deduce that isomerization is not the rate-limiting step. An analysis of the lineshapes suggested that the unbinding rate (koff) was about 45 /s, which is close enough to the catalytic rate to indicate that this step is rate-limiting.

But if koff is rate-limiting for this reaction, and the koff for R55A and R55K is dramatically increased (as it must be, with lower affinity), we ought to be seeing a higher rate in the ZZ-exchange experiments, or maybe even not seeing independent cross-peaks at all. How can this reaction be going slowly enough to be seen by this technique? As it turns out, the titrations hold the key. When CAN is saturated with R55A CypA, you can clearly see independent cis and trans peaks in the bound state (Figure 6, partially reproduced at right). This means that cis-trans interconversion on the enzyme has gotten much slower.

In fact, the presence of those two peaks means that we can use the ZZ-exchange experiment again, this time to determine the on-enzyme interconversion rate directly. The answer we get is about 20 /s, which is, within error, equivalent to the rate of the full cycle for this mutant. That means the rate-limiting step is no longer the unbinding of substrate, but rather the isomerization step itself. There’s only a minor change in overall catalytic efficiency, but this is the result of large changes in the rates of the individual steps that happen to cancel each other out.

Implications for the study of CypA-associated pathways

The best evidence available at the time supported the decisions Saphire et al. made in setting up their experiment. Previous work had clearly shown that an H126Q mutation of CypA significantly reduces the protein’s incorporation into virions (4,7). Saphire et al. made an H126A mutation on this basis and seemingly assumed that the activity would be similar (6). Unfortunately, the evidence from the NMR spectra is that H126A binds to the capsid protein perfectly well and also catalyzes its isomerization. This does not prove that the catalytic function of CypA is important for HIV-1 infectivity. However, on the basis of the existing experiments that possibility cannot yet be ruled out.

More broadly, these results demonstrate that claims about CypA’s role in biology cannot be based on mutant studies alone. The mutants discussed here alter, rather than abolish, CypA’s catalytic activity towards a biological substrate. Even those that appear not to bind the substrate in certain assays still display catalysis, because the strength of binding that is required for successful catalysis is considerably less than what is required for, say, a successful co-precipitation. The standard for assessing how a mutation has changed an enzyme’s behavior needs to be one that pays attention to the various steps of the reaction and how changes in particular rates can compensate one another. Experiments that rely on overexpression of a CypA mutant are particularly vulnerable to erroneous interpretation, because adding more enzyme is always an efficient way to compensate for a loss of activity and binding.

CypA and its related domains are very highly conserved across all vertebrates, yet its function was preserved even when apparently critical residues were dramatically altered by mutation. Our existing knowledge of protein sequences is limited to just a few examples from a relatively tiny number of species, and our structural and biochemical data encompass just a fraction of that. Assessments based on these databases are likely to underestimate the functionally viable sequence space. Descriptions of function based on model systems are also suspect. That goes for this system too — the findings about CypA activity made with respect to CAN are not necessarily any more generalizable than the chymotrypsin assay results. CypA can bind an enormous number of potential targets, and what is true for one may not be true for another. Whenever possible, catalytic activity and binding affinity ought to be verified directly on the substrate of interest. Otherwise, you might find that an enzyme you thought was dead is still stumbling along.


1) Bosco, D.A., Eisenmesser, E.Z., Clarkson, M.W., Wolf-Watz, M., Labeikovsky, W., Millet, O., & Kern, D. (2010). “Dissecting the Microscopic Steps of the Cyclophilin A Enzymatic Cycle on the biological substrate HIV-capsid by NMR” Journal of Molecular Biology DOI: 10.1016/j.jmb.2010.08.001

2) Bosco, D.A., Eisenmesser, E.Z., Pochapsky, S., Sunquist, W.I., and Kern, D. (2002) “Catalysis of cis/trans isomerization in native HIV-1 capsid by human cyclophilin A.” Proceedings of the National Academy of Sciences, 99(8), 5247-5252. DOI: 10.1073/pnas.082100499

3) Braaten, D., & Luban, J. (2001). “Cyclophilin A regulates HIV-1 infectivity, as demonstrated by gene targeting in human T cells” The EMBO Journal, 20 (6), 1300-1309 DOI: 10.1093/emboj/20.6.1300 OPEN ACCESS

4) Dorfman, T., Weimann, A., Borsetti, A., Walsh, C.T., & Göttlinger, H.G. (1997). “Active-site residues of cyclophilin A are crucial for its incorporation into human immunodeficiency virus type 1 virions.” Journal of Virology, 71 (9), 7110-3 PMCID: PMC192007 OPEN ACCESS

5) Gamble, T.R., Vajdos, F.F., Yoo, S., Worthylake, D.K., Houseweart, M., Sundquist, W.I., & Hill, C.P. (1996). “Crystal Structure of Human Cyclophilin A Bound to the Amino-Terminal Domain of HIV-1 Capsid” Cell, 87 (7), 1285-1294 DOI: 10.1016/S0092-8674(00)81823-1 OPEN ACCESS

6) Saphire, A.C.S., Bobardt, M.D., & Gallay, P.A. (2002). “trans-Complementation Rescue of Cyclophilin A-Deficient Viruses Reveals that the Requirement for Cyclophilin A in Human Immunodeficiency Virus Type 1 Replication Is Independent of Its Isomerase Activity” Journal of Virology, 76 (5), 2255-2262 DOI: 10.1128/jvi.76.5.2255-2262.2002 OPEN ACCESS

7) Zydowsky, L., Etzkorn, F., Chang, H., Ferguson, S., Stolz, L., Ho, S., & Walsh, C. (1992). “Active site mutants of human cyclophilin A separate peptidyl-prolyl isomerase activity from cyclosporin A binding and calcineurin inhibition” Protein Science, 1 (9), 1092-1099 PMCID: PMC2142182 OPEN ACCESS

Mar 232010
ResearchBlogging.orgA protein has several different levels of structure. The primary structure is the arrangements of atoms and bonds, and it is formed in the ribosome by the assembly of amino acids as directed by an RNA template. The secondary structure is the local topology, the helices and strands, and this forms mostly because of the release of energy through the formation of hydrogen bonds. The tertiary structure is the actual fold of the protein, the way helices, strands, and loops are arranged in space. The fold forms primarily because of the favorable entropy of burying the protein’s hydrophobic groups where water cannot access them, analogous to the formation of an oil droplet in water. This suggests that, in addition to the well-known phenomenon of proteins denaturing, or losing their higher-order structure, under conditions of high heat, proteins might also denature when they get too cold.

As you might remember from your chemistry classes, the change in free energy due to a reaction under conditions of constant pressure is given by:
ΔG = ΔH – T ΔS
Where ΔH is the change in enthalpy (i.e. the heat released or absorbed by a reaction), ΔS is the change in entropy, and T is the temperature of the system in Kelvin. Here, the change we are talking about is the transition from the folded state to some unfolded state. Simplistically, since the entropic contribution is scaled by the temperature, one can imagine that for a reaction with favorable entropy and unfavorable enthalpy, lowering the temperature could cause the reaction to reverse. Protein folding is only marginally favorable at biological temperatures, so one could easily imagine that lowering the temperature enough could cause a protein to prefer the unfolded state.

Of course, this is an oversimplification: the entropy and enthalpy of a particular protein state do not remain constant over all temperatures. Rather, they vary in a way determined by the heat capacity (Cp), such that ΔG as a function of temperature is (1):

ΔG(T) = ΔH(Tr) + ΔCp(T-Tr) – T [ΔS(Tr) + ΔCp ln(T/Tr)]
Where Tr is some reference state at which the thermodynamic parameters have been determined, and ΔCp is defined with respect to the native (folded) state. Because the various states of the protein have different Cp (unfolded chains typically have higher Cp), at certain temperatures above and below the biological optimum we can expect proteins to lose their higher levels of structure. Even this is still an oversimplification, of course, because it does not directly account for changes in water structure and cosolute properties with temperature. These features may cause ΔCp itself to vary with temperature rather than remain constant.

Unfortunately, for most proteins the temperature that favors unfolding lies below the freezing point of water, which makes this phenomenon difficult to study unless you do something unusual to your system. In 2004, Babu et al. (1) reported results from experiments that used reverse micelles to study the denaturation of ubiquitin at temperatures below freezing. By encapsulating a protein-water droplet in inverted micelles dissolved in pentane, it was possible to reduce the temperature to 243 K without causing freezing. These micelles also had the convenient property of tumbling quickly in the pentane, which allowed for reasonable NMR spectra even at these low temperatures. The appearance of the spectra they obtained indicated that the protein underwent a slow unfolding process with many different unfolded states, and also that the protein did not unfold in a cooperative fashion. Rather, it appeared that one contiguous region of the protein unfolded while the rest remained folded (the main helix was particularly stable).

This wasn’t expected, because ubiquitin apparently unfolds in a completely two-state manner when overheated. This being the case, what’s expected is for the protein to either be all folded or all unfolded, not some mixture of the two. However, cold does not affect all intramolecular contacts the same way. Lowering the temperature is expected to make hydrophobic interactions less favorable while not significantly affecting polar interactions like hydrogen bonds. This being the case, one might expect an α-helix to persist through a cold-denaturation transition, as happens in this case.

Something similar is observed in an upcoming paper in JACS from the Raleigh and Eliezer Labs (2), which approaches cold denaturation using a mutant form of the C-terminal domain of ribosomal protein L9. An isoleucine to alanine mutation at residue 98 of this domain doesn’t appear to significantly alter the structure, but it causes the protein to denature somewhere in the high teens. At 12 °C the unfolded state is about 80% of the visible population, and this is where Shan et al. performed their NMR experiments. They assigned the unfolded state using standard techniques and then decided to see what they could learn from the chemical shifts.

As I’ve mentioned before, the chemical shift of a nucleus depends on the probability distribution of the surrounding electrons, and therefore is sensitive to the strength, composition, and angles of the atom’s chemical bonds. Because the dihedral angles of the protein backbone are a good proxy for the secondary structure, one can use the chemical shifts of particular atoms to determine whether a given residue is in a helix or strand. When they performed this analysis, Shan et al. noticed two major differences between the native and cold-denatured states of the protein. The first was that the helix and strand propensities of the denatured protein were much lower than the folded form, as expected. In addition, however, they noticed that one loop of the protein had gained α-helical character. That is, it seemed like an α-helix had actually gotten longer as a result of the unfolding.

This doesn’t mean that denaturing the protein added secondary structure. The low values in the output from the algorithm Shan et al. used suggest that the secondary structure in this denatured state forms only transiently. However, the chemical shifts suggest, and other structural data appear to confirm, that this region of the protein has an increased propensity to inhabit a helical structure as a consequence of the unfolding.

These results emphasize the fact that the “unfolded state” isn’t as simple as it’s often described. Residual structure persists in unfolded states of many proteins, and unfolded ensembles of one protein generated through different means (heat, cold, pH, cosolutes) may not resemble each other. Unlike unfolding at high temperature, cold denaturation of ubiquitin appears to be non-cooperative. In both ubiquitin and L9, it appears that helices are robust to the unfolding process, persisting and even propagating as the protein denatures. While some of these features may be held in common between different kinds of denatured states, others may be unique to particular denaturation conditions. The lingering question is which of these unfolded ensembles best resembles the denatured state that exists under biological conditions, giving rise to misfolded states and their associated diseases.

(1) Babu, C., Hilser, V., & Wand, A. (2004). Direct access to the cooperative substructure of proteins and the protein ensemble via cold denaturation Nature Structural & Molecular Biology, 11 (4), 352-357 DOI: 10.1038/nsmb739

(2) Shan, B., McClendon, S., Rospigliosi, C., Eliezer, D., & Raleigh, D. (2010). The Cold Denatured State of the C-terminal Domain of Protein L9 Is Compact and Contains Both Native and Non-native Structure Journal of the American Chemical Society DOI: 10.1021/ja908104s

 Posted by at 10:00 PM
Feb 262010
ResearchBlogging.orgSpongiform encephalopathies are transmissible diseases that can have a major economic impact on agricultural exports, and pose a significant challenge for surveillance of the food supply. Scientists generally believe that these diseases are transmitted via a self-propagating, aberrant conformation of the prion protein (PrP). This prion hypothesis suggests that PrP alone should be sufficient to cause symptoms or death. If this hypothesis is true, then it should be possible to reproduce the disease using recombinant proteins expressed in yeast or bacteria. In tomorrow’s Science, researchers from Columbus and Shanghai report that they have managed to do this, establishing that PrP alone can account for prion disease transmission.

Previously, other groups had successfully produced recombinant PrP (recPrP) and generated amyloid fibrils that appeared to contain the pathogenic conformation (PrPSc). When injected into mice, however, these amyloids had limited infectivity, which raised doubts as to whether these fibers are the cause of the disease. Wang et al. decided to take a different route, using a technique known as protein misfolding cyclic amplification (PMCA). In this approach, misfolded aggregates of a protein are broken up using sound waves and then incubated with normal, folded protein. If the misfolded protein can cause normal protein to adopt an aberrant conformation (as PrPSc can), then the misfolded protein will be amplified. By performing many cycles of this experiment, one can in principle produce a very large amount of PrPSc from a single misfolded chain.

Wang et al. also added some ingredients to their reactions that they believed would promote prion formation: RNA and a lipid called POPG. Under these conditions, they detected a protease-resistant protein after 17 rounds of PMCA amplification. Under normal circumstances, PrP is cleaved by the protease, like a pair of scissors cutting a string. If PrP has aggregated, however, it becomes much more difficult to cut, as if you were using safety scissors to cut a thick hemp rope. The researchers discovered that the protein from this reaction (which they called rPrP-res) could cause normal protein to also become protease resistant. And, when they treated protein from mouse brains with rPrP-res, they found that it, too, formed protease-resistant aggregates.

To really put the hypothesis to the test, however, tests in live animals were needed. The researchers injected one group of animals with the product of a PMCA seeded with rPrP-res. They also injected three additional groups of animals with control cocktails to prove that neither the non-protein ingredients, nor the unprocessed protein, caused the disease. None of the control animals developed symptoms of encephalopathy during the experience, but all of the mice injected with the PMCA product died in about five months, while displaying clinical signs of prion disease. Their brains were inspected post-mortem using histological and molecular means. The brains clearly showed the formation of the vacuoles that give spongiform encephalopathy its name. Additionally, protease-resistant PrP was detected in homogenates of the brain tissue, indicating that the rPrP-res had propagated in the living mouse brains, causing disease and eventual death. When these homogenates were injected into the brains of other healthy mice, a similar pattern of pathology recurred. This proved that the effects of rPrP-res could be serially propagated, just as prion disease is.

These results leave little room for doubt that misfolded PrP is sufficient to cause prion disease; no other infectious agent was required. The effectiveness of RNA and POPG in promoting the pathogenic conformation may indicate that these or similar molecules play a role in the spontaneous development of prion disease. Aside from adding to our knowledge directly, this research has the potential to significantly increase our ability to investigate prion disease. The ability to produce bona fide infective prion molecules in vitro from recombinant protein opens up new avenues for experiments in structural biology and biochemistry that may enable us to cure or entirely prevent these diseases, rather than just trying to contain them.

Wang, F., Wang, X., Yuan, C., & Ma, J. (2010). Generating a Prion with Bacterially Expressed Recombinant Prion Protein Science, 327 (5969), 1132-1135 DOI: 10.1126/science.1183748