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PROTEIN | Interactions and Reactions Involved in Food Processing
Decomposition of Amino Acids
Studies of model systems have revealed that protein cleavage and degradation of side-chains, rather than the formation of protein networks, are the preferred reactions when the water content of protein–lipid mixtures decreases. Examples for the extent of losses in amino acids in a protein in the presence of an oxidized lipid are presented in Table 3 . The strong dependence of this loss on the nature of the protein and on reaction conditions is obvious. Degradation products obtained in model systems of pure amino acids and oxidizing lipids are listed in Table 4 .
Table 3 . Amino acid losses occurring in proteins exposed to peroxidized lipids
|Reaction system||Reaction conditions||Amino acids lost a (% loss)|
|Casein||Linoleic acid ethyl ester||24 h||55 b||Lys(10), Thr(10), Val(10), Ala(9), Tyr(8), Phe(8), Ser(8), Arg(8), Asp(8) c,d|
|Casein||Linoleic acid ethyl ester||4 days||60 e||Lys(50), Met(47), Ile(30), Phe(30), Arg(29), Asp(29), Gly(29), His(28), Thr(27), Ala(27), Tyr(27) c,d|
|Ovalbumin||Linoleic acid ethyl ester||24 h||55 b||Met(17), Ser(10), Lys(9), Ala(8), Leu(8) c,d|
|Ovalbumin||Linoleic acid ethyl ester||4 days||60 e||Lys(50), Met(42), Leu(22), His(21), Val(21) c,d|
|Ribonuclease||Linoleic acid||40 min||37 b||Met(99), Tyr(62), His(54), Lys(51), Cys(40) c|
|Trypsin||Linoleic acid||40 min||37 b||Met(83), His(12) c|
|Lysozyme||Linoleic acid||8 days||37 b||Trp(56), His(42), Lys(17), Met(14), Arg(9)|
Data from Gardner HW (1979) Lipid hydroperoxide reactivity with proteins and amino acids: a review. Journal of Agricultural and Food Chemistry 27: 220–229 and references therein.
a Only the most labile amino acids are listed. b Aqueous system. c Trp not analyzed. d Cys not analyzed. e 80% relative humidity.
Table 4 . Products formed from amino acids exposed to peroxidizing lipids
|Reaction system||Compounds formed from amino acids|
|Histidine||Methyl linoleate||3-(Imidazol-4-yl)lactic acid, 3-(imidazol-4-yl)acetic acid histamine, valine, aspartic acid, ethylamine|
|Cysteine||Ethyl arachidonate||Cystine, hydrogen sulfide, alanine|
|Linoleic acid||Cystine, cysteic acid, cystine disulfoxide|
|Methionine||Methyl linoleate||Methionine sulfoxide|
|Lysine||Methyl linoleate||Diaminopentane, aspartic acid, glycine, alanine, α-aminoadipic acid, pipecolinic acid, 1,10-diamino-1,10-dicarboxydecane|
Modified from Gardner HW (1979) Lipid hydroperoxide reactivity with proteins and amino acids: a review. Journal of Agricultural and Food Chemistry 27: 220–229.
OMA1 and YME1L differentially modulate OPA1 fusion activity
Whereas human OPA1 has eight different mRNA splice variants (Supplemental Figure S1A), mouse tissues express only four–-isoforms 1, 5, 7, and 8 (Akepati et al., 2008) (Figure 1A). Each of isoforms 1 and 7 produces a mixture of l-OPA1 and one or two versions of s-OPA1, respectively. The short isoforms are produced by proteolytic cleavage of the long isoform by the mitochondrial inner membrane proteases OMA1 (at the S1 site) and YME1L (at the S2 site). In contrast, proteolytic processing of isoforms 5 and 8 goes to completion and results in only short isoforms (Song et al., 2007).
FIGURE 1: OMA1 and YME1L differentially regulate OPA1 fusion activity. (A) Schematic of mouse OPA1 protein isoforms, showing the origin of protein bands a–e. MEFs express isoforms 1, 5, 7, and 8. Isoforms 1 and 7 produce long forms that constitute bands a and b and short forms that constitute bands c–e. Isoforms 5 and 8 produce exclusively short forms expected to comigrate with bands c–e, but for simplicity, bands c–e are labeled according to their origin from isoforms 1 and 7. The MPP (mitochondrial processing peptidase), S1 (by OMA1), and S2 (by YME1L) cleavage sites are indicated with arrows. The orange line and arrows show that bands c and d are derived from long isoform 7/a, and the pink line and arrow show that band e is derived from the long isoform 1/b. (B) Western blot analysis of OPA1 bands in Oma1 and Yme1l mutant MEFs. Five OPA1 bands (a–e) are apparent in wild-type MEFs. Oma1-null MEFs lack c and e Yme1l-null MEFs lack d and Oma1/Yme1l-null MEFs lack all the short forms, c–e. Tubulin was used as loading control. (C) Representative images of mitochondrial morphology (mito-DsRed) in WT and protease-null MEFs in different media. GLY: high glucose medium OXI: OXPHOS-inducing medium CHX: high glucose medium with 10 μM cycloheximide. Insets show magnified view. Scale bar, 5 μm. (D) Quantification of mitochondrial morphology of cells in C. In each experiment, 100 cells were scored. Error bars show SD from three independent experiments. (E) Comparison of mitochondrial fusion rates in vivo in different media between WT and Oma1/Yme1l-null MEFs. Fusion activity was measured by the intensity reduction of PA-GFP as a function of time. Error bars represent SDs from at least six independent measurements.
In mouse embryonic fibroblasts (MEFs), this group of four mRNA isoforms gives rise to five protein bands when cell lysates are resolved on SDS–PAGE (labeled a–e in Figure 1, A and B). Bands a and b are the long forms arising from isoform 7 and isoform 1, respectively (Figure 1A). Bands c and d are S1- and S2-cleavage products of isoform 7 generated by OMA1 and YME1L, respectively. Band e is the short, S1-cleaved form of isoform 1. Isoforms 5 and 8 generate only short isoforms that would be expected to contribute to bands c–e (Song et al., 2007). The same set of OPA1 bands has been observed in other studies, although the relative intensities can vary, perhaps due to differences in culture conditions (Anand et al., 2014).
The dependencies of bands c–e on the OMA1 or YME1L proteases were confirmed by the OPA1 band pattern in lysates from Oma1-null or Yme1L-null cells (Figure 1B). The OPA1 band patterns observed in the protease mutant cells are tabulated in Supplemental Figure S1B. The identities of the Oma1-null and Yme1L-null cells were confirmed by Western blotting against the relevant proteases (Supplemental Figure S1C). Notably, there is a longer band emerging in both the Yme1l-null and Yme1l/Oma1-null cells, likely due to the disruption of YME1L-dependent short isoform formation (i.e., isoform 8).
To understand the role of these proteases in regulating OPA1 fusogenic function, we examined Oma1/Yme1l double knockout MEFs, which contain only long forms of OPA1 (Anand et al., 2014). In standard high glucose culture medium, these cells show increased mitochondrial fragmentation compared with wild-type (WT) cells (Figure 1, C and D). Many Oma1/Yme1l mutant cells nevertheless show short tubular mitochondria (Figure 1D), so the degree of mitochondrial fragmentation is substantially less severe than in Mfn1/Mfn2-null or Opa1-null cells (Chen et al., 2005, 2007). Oma1-null MEFs show normal mitochondrial morphology profiles, and Yme1l-null MEFs show highly fragmented mitochondria (Figure 1D). The overall trends–that mitochondria in Oma1/Yme1l mutant cells are more fragmented than in WT cells, but less so than in Yme1l mutant cells–are consistent with a previous study (Anand et al., 2014). OPA1 band patterns are similar under the different media conditions (Supplemental Figure S1D).
We tested how these mutant cells respond to culture conditions that increase the level of mitochondrial fusion. Cultured MEFs increase mitochondrial fusion activity and show mitochondrial elongation when cultured in medium that induces oxidative phosphorylation (OXPHOS). This response has been shown to depend on YME1L activity (Mishra et al., 2014). Consistent with this idea, we found mitochondrial elongation caused by OXPHOS-inducing medium to be abrogated in Yme1l-null and Oma1/Yme1l-null MEFs (Figure 1, C and D). As measured in the photoactivatable (PA)–green fluorescent protein (GFP) fusion assay, mitochondrial fusion under basal conditions was substantially reduced in Oma1/Yme1-null MEFs compared with that in WT cells and did not increase in response to OXPHOS-inducing medium (Figure 1E). In contrast, cells of all genotypes tested in this study show robust mitochondrial elongation in response to cycloheximide (CHX) treatment, which causes stress-induced mitochondrial hyperfusion (Tondera et al., 2009) (Figure 1, C–E). Hyperfusion is induced rapidly after the addition of CHX (Figure 1E). CHX treatment enhanced PA-GFP signal dilution in both WT and Oma1/Yme1l-null cells. WT cells showed a substantially greater fusion rate than the latter, indicating the reduced fusion capability in Oma1/Yme1l-null cells even in the presence of a strong fusion stimulus. Taken together, our observations agree with the previous finding that Oma1/Yme1l-null cells, containing only l-OPA1, retain mitochondrial fusion activity (Anand et al., 2014). However, the level of fusion activity is substantially lower compared with that in WT cells. The OMA1 and YME1L proteases differentially regulate OPA1 function. Stress-induced hyperfusion does not depend on either protease, consistent with the report that l-OPA1 is sufficient in mediating fusion under this stress condition (Tondera et al., 2009). In contrast, OXPHOS-induced mitochondrial elongation is dependent on YME1L activity, as noted previously (Mishra et al., 2014).
The Opa1 S2 site is necessary for mediating OXPHOS-induced fusion in vivo
To clarify the functions of YME1L-dependent cleavage of OPA1, we used CRISPR-Cas9–mediated gene targeting in MEFs to generate mutations in the Opa1 genomic locus at exon 5b, which encodes the S2 cleavage site (Figure 2, A and B). We hoped to generate deletions in exon 5b that would disrupt the Opa1 reading frame, resulting in a downstream premature stop codon that would prevent the formation of functional OPA1 isoforms containing S2. In this scenario, the mutant cells would be expected to have only long isoform 1 (band b) and the corresponding S1 cleavage product (band e). Bands a, c, and d would be missing, because they arise from exon 5b–containing transcripts (Figure 2B).
FIGURE 2: The S2 site encoded by Opa1 exon 5b is necessary for OXPHOS-induced fusion in vivo. (A) Schematic of exon 5b, showing locations of S2 and the CRISPR-Cas9 gRNA target. Deletions present in clones 1 and 2 are indicated. (B) Schematic of OPA1 isoform composition after exon 5b–containing isoforms are eliminated Red Xs indicate isoforms expected to be missing in mutant cells. (C) Western blot analysis of ∆exon 5b MEF clones. Clones 1 and 2 are positive clones that show the expected disappearance of bands a, c, and d. The last lane is a negative clone. Tubulin was used as loading control. (D) Quantification of mitochondrial morphology in WT and two ∆exon 5b clones in the indicated media. One hundred cells were counted for each experiment. Error bars indicate SD from three independent experiments.
After transient expression of Cas9 and gRNA (guide RNA) against the exon 5b target in MEFs, we identified multiple colonies that lacked bands a, c, and d (Figure 2C). To ensure against clone-specific effects, we used three or more ∆exon 5b clones in all our experiments, and results from two different clones are shown in the figures. DNA sequencing showed that clone 1 has a homozygous two-nucleotide deletion and clone 2 has two longer deletions each causing frameshifting (Figure 2A).
The ∆exon 5b cells showed normal mitochondrial profiles when grown in regular glucose-containing medium. In addition, they showed a normal stress-induced hyperfusion response, dramatically elongating mitochondria upon CHX stimulation (Figure 2D and Supplemental Figure S2A). In OXPHOS-inducing medium, however, the mutant cells did not show any elongation of mitochondria compared with growth in control medium, indicating that S2-cleavage of Opa1 is required for OXPHOS-induced hyperfusion. Although OXPHOS-inducing medium does not promote mitochondrial elongation in ∆exon 5b cells, we did not detect physiological consequences. OXPHOS-inducing medium caused upregulation of oxygen consumption in these cells (Supplemental Figure S2B), indicating that the increase in respiratory function is not dependent on mitochondrial elongation. Moreover, both WT cells and ∆exon 5b cells showed similar cell growth after switching to OXPHOS-inducing medium (Supplemental Figure S2C). In addition to MEFs, we generated two ∆exon 5b mouse lines. Western analysis of mutant tissues showed abrogation of Opa1 exon 5b–containing protein bands (Supplemental Figure S2D). Despite this biochemical change in OPA1 isoforms, the mutant mice showed no obvious physiological dysfunction and gained weight normally (Supplemental Figure S2E) for at least 1 yr of age.
Detection of a new OPA1 cleavage site in WT and ∆exon 5b cells
In the course of this study, we switched to a higher-resolution gel system to study these OPA1 protein isoforms. With this new system, what was previously designated as band d in WT cells could be resolved into doublet bands, which we designate as d and d′ (Figure 3A). Upon reanalysis, we realized that ∆exon 5b cells produce two short Opa1 bands, d′ and e. With deletion of the YME1L-dependent S2 site, ∆exon 5b cells would be expected to show only OMA1-mediated processing of OPA1. In ∆exon 5b/Oma1-null cells, band e is missing but d′ band remains, indicating that it is OMA1-independent (Figure 3A).
FIGURE 3: Detection of a new OPA1 cleavage in WT and ∆exon 5b cells. (A) Higher-resolution Western analysis of WT, ∆exon 5b, and ∆exon 5b/Oma1-null MEFs. Note the additional band located under d in WT MEFs. This band (referred to as d′) is more clearly seen in ∆exon 5b and ∆exon 5b/Oma1-null clones. HSP60 was used as loading control. (B) Dependence of band d′ on Yme1l. Cells expressing the indicated shRNA were analyzed by Western blotting. The two sets of panels were run on the same gel. HSP60 was used as loading control. (C) PCR analysis of Opa1 transcripts in ∆exon 5b cells. The forward primer is located in exon 3, and the reverse primer is located in exon 7 (Supplemental Figure S3C). Bands corresponding to isoforms 1 and 5 were confirmed by sequencing. (D) Schematic of full-length OPA1 isoforms 1 and 5. (E) Western blot analysis of the short form produced by isoform 5. HSP60 was used as loading control. (F) Dependence of isoform 5 band d′ on YME1L. FLAG-tagged human OPA1 (isoform 1 or 5) was expressed in cells of the indicated genotype and analyzed by Western blotting against FLAG. HSP60 was used as loading control.
Because YME1L is the other known intermembrane protease involved in OPA1 posttranslational processing, we used short hairpin RNA (shRNA) to test whether it is essential for the production of band d′. The knockdown efficiencies of the Yme1l and Oma1 shRNAs were confirmed by Western blotting (Supplemental Figure S3, A and B). In WT cells, knockdown of YME1L caused the disappearance of bands d and d′, whereas knockdown of OMA1 caused the disappearance of bands c and e (Figure 3B). The dependence of d′ on YME1L but not OMA1 was further confirmed in both Δexon 5b/Oma1-null cells and Δexon 5b cells (Figure 3B).
To identify the source of the new short OPA1 band d′, we used PCR analysis of cDNA from Δexon 5b cells to identify the major remaining Opa1 mRNA transcripts. Using primers flanking the alternative splicing exons 4, 4b, and 5b to distinguish individual mRNA transcripts (Supplemental Figure S3C), we found that the mutant cells expressed mRNA splice variants 1 and 5 (confirmed by both PCR analysis and sequencing), with isoform 1 being slightly more abundant (Figure 3, C and D). It is not surprising that isoforms 7 and 8 are missing, because they contain exon 5b. Isoform 5 instead contains the alternative exon 4b (Figure 3D), and like other exon 4b–containing isoforms, was previously shown to be constitutively cleaved into a short isoform (Song et al., 2007). The constitutive cleavage of isoform 5 was assumed to occur at the S1 site, the only known cleavage site in this isoform (Song et al., 2007). However, we wondered whether this assumption might be incorrect, because cleavage of isoform 5 at S1 would lead to band e, not d′. To test whether isoform 5 could generate band d′, we expressed isoform 5 in Opa1-null cells. Isoform 5 generated a short isoform that was clearly distinct from e and comigrated with the d′ bands present in ∆exon 5b and ∆exon5b/Oma1-null cells (Figure 3E). This observation indicates that isoform 5 is the likely source of band d′. Its cleavage site is N-terminal to S1, as indicated by the slightly larger size of d′ compared with e.
To determine the protease responsible for cleavage of isoform 5 and further confirm that the YME1L-dependent band d′ indeed arises from isoform 5, we compared the behavior of C-terminal FLAG-tagged human isoforms 5 and 1 in mutant MEFs lacking OMA1 or YME1L. Consistent with previous studies showing cleavage at S1 by OMA1, isoform 1-FLAG produces a short isoform that is dependent on OMA1 (Figure 3F). When YME1L is absent, more of this short isoform is produced, presumably due to up-regulation of OMA1 activity (Wai et al., 2015). As expected, isoform 5-FLAG is completely processed in control MEFs. The absence of OMA1 does not affect the cleavage pattern, indicating that the production of isoform 5 short form is independent of OMA1 (Figure 3F). The removal of Yme1l completely abrogates production of this short form and results in a lower band that comigrates with the short isoform generated from isoform 1 (Figure 3F). In addition, a small amount of the long form, previously absent, is formed. These results indicate that, under normal conditions, isoform 5 is processed to completion at a site upstream of S1, in a YME1L-dependent manner, to produce band d′. When YME1L is absent, cleavage is activated at the downstream S1 site. Only the long form is generated from isoform 5 when both OMA1 and YME1L are absent (Figure 3F, last lane). Together, the results suggest that OPA1 isoform 5 is highly susceptible to protease processing, normally by YME1L. In the absence of YME1L, OMA1 processes isoform 5 into S1-cleaved OPA1. No additional proteases are sufficient for isoform 5 cleavage, as indicated by the formation of long isoform 5 when both Yme1l and Oma1 are absent.
Identification of a leucine-rich, YME1L-dependent cleavage site in exon 4b
Our results so far suggest that isoform 5 contains a novel YME1L-dependent cleavage site upstream of S1. To pinpoint the new cleavage site (which we designate as S3), we asked whether cleavage is dependent on exon 4b, the first exon upstream of exon 5. Knockdown of exon 4b with shRNA substantially reduced the intensity of band d′ in WT, Δexon 5b, and Δexon 5b/Oma1-null cells (Figure 4A, band d′ marked with asterisks). This dependence suggests that the cleavage site likely resides in exon 4b. To test this idea, we analyzed the behavior of a series of isoform 5 mutants (Mut 1–9) containing various short deletions within exon 4b (Figure 4B). Mutants 1, 2, and 3, containing deletions in the 5′ half of exon 4b, show no defect in isoform 5 cleavage processing, indicating that the first half of exon 4b does not contain the cleavage site (Figure 4C). Mutations 4, 5, and 6 result in small amounts of the long form, which normally is absent. In addition, these mutants show prominent production of band e. Mutants 7 and 8 also show the presence of the long form and band e. In addition, these mutants show novel cleavage products forming a smear above band e (Figure 4C). Mutation 9, which deleted the bridging amino acids between exons 4b and 5, generates band e and produces new cleavage products without generating a long form of isoform 5. These results suggest that S3 cleavage is sensitive to mutations in the 3′ half of exon 4b, and particularly to mutations (mutants 7 and 8) affecting a stretch of five consecutive leucines (amino acids 199–203 in isoform 5). Interestingly, the most prominent effects of the mutations are in the appearance of the long form and novel cleavage products, rather than in elimination of band d′.
FIGURE 4: Identification of the YME1L-dependent OPA1 S3 cleavage site within exon 4b. (A) Dependence of band d′ on exon 4b. shRNA against Opa1 exon 4b was expressed in cells of the indicated genotype. Asterisks highlight band d′. HSP60 was used as loading control. (B) Schematic of Opa1 exon 4b mutants. At the top, the polypeptide sequences encoded by exon 4b (red) and exon 5 (blue) are shown. The red arrow indicates the known S1 cleavage site in exon 5 the green arrows indicate the new S3 sites found in this study. The isoform 5 mutants are listed, along with amino acids deleted within exon 4b. (C) Effect of exon 4b mutants on OPA1 processing. FLAG-tagged exon 4b mutants of human isoform 5 were expressed in Opa1-null MEF cells and analyzed by Western blot. FLAG-tagged WT isoforms 1 and 5 were used as controls. HSP60 was used as loading control. (D) Effect of OMA1 and YME1L on OPA1 processing in isoform 5 mutants. WT human Iso 5 and its mutants 2, 6, and 7 were expressed in Oma1- and Yme1l- null MEFs, and FLAG-tagged OPA1 band patterns were analyzed by Western blot. HSP60 was used as loading control. (E) Identification of likely S3 cleavage sites within exon 4b by tandem mass spectrometry of band d′. Band d′ was produced by expression of FLAG-tagged Opa1 isoform 5 in 293T cells, purified by anti-FLAG immunoprecipitation, resolved by SDS–PAGE, digested by trypsin, and subjected to tandem mass spectrometry. Left, Schematic indicates the three ragged N-terminal peptides identified, colored according to their relative intensity. Right, Plot of the intensity of the three peptides. (F) Effect of penta-leucine mutants on generation of band d′. As indicated, alanines were used to replace 1, 2, or 3 leucines within the C-terminal penta-leucine stretch encoded by exon 4b. FLAG-tagged mutants of isoform 5 were expressed in Opa1-null cells and analyzed by Western blot. HSP60 was used as loading control. (G) Schematic of the OPA1 isoform profile in ∆exon 5b MEFs. Green arrow represents the new S3 site identified.
To determine the proteases responsible for cleavage of the mutants, we expressed the FLAG-tagged mutants in protease mutant MEFs (Figure 4D). In mutants 6 and 7, the smallest cleavage product disappeared in Oma1-null cells, indicating that it is OMA1-dependent. In mutant 7, there are larger short forms that are YME1-dependent. In the absence of both OMA1 and YME1L, no cleavage was observed, demonstrating that these two proteases are essential for isoform 5 processing.
To identify the cleavage site more precisely, we used tandem mass spectrometry to determine the N-terminal sequence of band d′, produced by overexpression of FLAG-tagged human isoform 5 in 293T cells. Band d′ was collected by immunoprecipitation against the FLAG tag, resolved on SDS–PAGE, and isolated for biochemical analysis. Peptides were generated by trypsin digestion and identified by liquid chromatography coupled with nanospray ionization tandem mass spectrometry (LC-MS). Of the peptides identified, only three had nontryptic N-termini. The two predominant ones had N-termini located within exon 4b, beginning with the second to last leucine and last leucine of the penta-leucine stretch (Figure 4E). The latter peptide had by far the highest intensity, suggesting that it represents the predominant S3 cleavage product. To test this idea, we designed isoform 5 mutants (mutants 10–13) in which the last 1–3 leucines of the penta-leucine stretch were substituted with alanine. We found that substitution of a single leucine (mutants 12 and 13) had little effect on band d′ production, though a small level of larger cleavage products could be observed (Figure 4F). When both of the last two leucines were substituted (mutant 11), there was a more substantial level of aberrant cleavage, and the long isoform appeared. Both these effects were increased when the last three leucines were substituted (mutant 10). Taken together, the LC-MS and mutational analysis strongly argue that the S3 cleavage site lies within the penta-leucine stretch encoded by the end of exon 4b. Although the product ion spectra indicated that cleavage primarily occurs N-terminal to the last leucine, the mutational phenotypes suggest that cleavage can occur at other locations very near the penta-leucine stretch (Figure 4C, lanes 9–13). None of the mutations completely eliminates production of band d′, suggesting that cleavage can still occur very close to S3 when the site is mutated. It is also apparent that isoform 5 is highly prone to proteolytic processing—when the S3 site is mutated, cleavage is activated at S1 or at cryptic sites N-terminal to S3, leading to variant cleaved forms of isoform 5. Our analysis indicates that ∆exon5b cells retain two dominant Opa1 mRNA transcripts, isoforms 1 and 5. This simplified transcript profile results in one l-OPA1 (from isoform 1) and two s-OPA1 (isoforms 1/S1 and 5/S3) (Figure 4G).
The S3-cleaved short form of isoform 5 regulates mitochondrial tubulation
In both humans and mice, half of the Opa1 splice forms contain exon 4b and are constitutively cleaved to yield exclusively short isoforms (Song et al., 2007). This raises the issue of why Opa1 encodes multiple mRNA splice forms whose only apparent purpose is to generate short isoforms. We used isoform 5 to address this issue. Consistent with previous studies (Song et al., 2007), isoform 5 has almost no mitochondrial elongation activity when expressed in Opa1-null MEFs, which have highly fragmented mitochondria due to the complete loss of inner membrane fusion (Figure 5A and Supplemental Figures S5A and S4A). Isoform 5 mutations that disrupt S3 processing, however, show substantial mitochondrial fusion activity, as indicated by their ability to elongate mitochondria (Figure 5A and Supplemental Figures S5A and S4, A and B). Because these mutants produce a variety of short forms as well as a long form, these results fit with the model that a mixture of long and short isoforms of OPA1 is important for mitochondrial fusion activity (Song et al., 2007). Alternatively, these results could be explained by proposing that it is the production of the long isoform that induces mitochondrial fusion activity.
FIGURE 5: The S3-cleaved short form of isoform 5 regulates mitochondrial morphology. (A) Elongation of mitochondria by isoform 5 mutants. Isoform 5 mutants were expressed in Opa1-null cells, and mitochondrial morphologies were quantified. Error bars indicate SD from six independent experiments. (B) Depletion of band d′ by shRNA against exon 4b and reexpression of d′ by isoform 5. Ectopic expression of isoform 5 is sufficient to overcome the shRNA effect and produce d′. OPA1 isoforms were analyzed by Western blotting against OPA1. HSP 60 was used as loading control. (C) Effect of exon 4b shRNA on mitochondrial morphology. Error bars indicate SD from three independent experiments. (D) Analysis of bicistronic OPA1 expression. Opa1-null cells expressing isoform 1 or isoform 1 plus isoform 5 were analyzed by Western blotting against OPA1. ∆exon 5b cells are shown for comparison. HSP60 was used as loading control. (E) Effect of band d′ on mitochondrial morphology. Quantification of mitochondrial morphology of Opa1-null MEFs expressing isoform 1 with YFP or isoform 1 with isoform 5. Error bars indicate SD from three independent experiments. P values for the fragmented, short tubular, long tubular, and interconnected were 4.8 × 10 –4 , 0.04, 0.001, and 9.9 × 10 –7 , respectively. P values were calculated by Student’s t test. (F) Model showing the OPA1 isoforms under normal and S3 disrupted conditions. When S3 is disrupted (bottom panel, box), cleavages at S1 and cryptic sites are activated, preserving production of s-OPA1. OM: outer membrane, IM: inner membrane.
To distinguish between these two models, we further probed the physiological function of the short form of isoform 5. We used shRNA to knock down exon 4b in Δexon 5b and Δexon 5b/Oma1-null cells, which have a simplified portfolio of Opa1 mRNA splice forms. As expected, knockdown of exon 4b resulted in reduction of band d′ in both cell lines (Figures 4A and 5B). In the Δexon 5b MEFs, exon 4b knockdown leads to dramatic mitochondrial fragmentation (Figure 5C and Supplemental Figure S5B). Similarly, in Δexon 5b/Oma1-null cells, knockdown of exon 4b results in mitochondrial fragmentation (Figure 5C and Supplemental Figure S5B). To verify that the knockdown result is not due to off-target effects, we reexpressed isoform 5 in the exon 4b knockdowns (Figure 5B) and observed elongation of mitochondrial morphology in both cell lines (Figure 5C and Supplemental Figure S5B).
To further test this idea that a short form of OPA1 can collaborate with a long form to regulate mitochondrial fusion, we developed an expression system to coexpress two different OPA1 splice forms in Opa1-null MEFs. We constructed a bicistronic expression vector to generate isoform 1 along with isoform 5, or isoform 1 along with control YFP. Western blot confirmed the expected OPA1 band pattern in these cells (Figure 5D). With isoform 1 expression alone, the long band b and the short band e were produced. With bicistronic expression of isoforms 1 and 5, an additional short form (d′ from isoform 5) was added. This dual expression recapitulated the bands observed in Δexon 5b cells (Figure 5D) and resulted in longer mitochondrial profiles. Significantly more cells contained interconnected mitochondria, and fewer cells had fragmented mitochondria, compared with the expression of isoform 1 alone (Figure 5E and Supplemental Figure S5C). These results suggest that S3-cleaved s-OPA1 can work with l-OPA1 to promote mitochondrial fusion. To determine whether this synergy requires GTP hydrolysis activity in s-OPA1, we used the bicistronic system to express wild-type isoform 1 with isoform 5 containing a dysfunctional GTPase domain (iso 5-G300E) in Opa1-null MEFs. The protein band pattern was similar in cells expressing Iso 1-IRES-Iso 5-G300E compared with cells expressing Iso 1-IRES-Iso 5 (Supplemental Figure S5D). The mutated isoform 5 was ineffective at promoting mitochondrial fusion in combination with wild-type isoform 1, demonstrating that GTPase activity is required in s-OPA1 to maintain fusion function (Supplemental Figure S5, C and E).
Specificity of Purified ADAM 10 for Peptide Substrates.
As a peptide substrate spanning the α-secretase cleaving site, we chose the octadecapeptide amide sequence, residues 11 in Aβ (numbering with respect of the N terminus of Aβ) (12). Its C terminus corresponds to the first extracellular residue of APP. After a 30-min incubation of Aβ(11) with ADAM 10, HPLC analysis showed cleavage of 28% of the starting peptide with two fragments arising (Table (Table1). 1 ). Analysis by electrospray ionization mass spectrometry identified them as the N-terminal hexapeptide ([M+H] + ion at m/z of 777.5) and the C-terminal dodecapeptide amide ([M+H] + ion at m/z of 2082.8) of the parent octadecapeptide amide. Thus ADAM 10 proteolytically cleaves between Lys-16 and Leu-17 as expected for an enzyme with α-secretase activity. After a 6-hr incubation, 69% of Aβ(11) were cleaved predominantly at this site with minor cleavage occurring after Leu-17 and Val-18 (Fig. (Fig.1 1 A and Table Table1). 1 ).
Specificity of purified ADAM 10 from bovine kidney for peptides spanning the cleavage site of shed proteins
|Protein||Sequence||% cleavage by bovine ADAM 10|
|30 min||6 hr|
|APP||EVHHQK ↓ LVFFAEDVGSNK-NH2||28||69|
|APP (A692G)||EVHHQK ↓ LVFFGEDVGSNK-NH2||14||54|
|IL-6 receptor||ANATSLPVQ ↓ DSSSV-NH2||0||0|
|Angiotensin-converting enzyme||TPNSAR ↓ SEGPLPDSGR-NH2||0||0|
|Pro-TNF-α||PLAQA ↓ VRSSSRTPSD-NH2||49||100|
Cleavage of peptides by ADAM 10 after 30 min or 6 hr was determined by HPLC analysis. ↓, proteolytic cleavage site.
Cleavage of peptides spanning the α-secretase cleavage site of APP by ADAM 10. (A) Peptide substrates were incubated in cleavage buffer in the absence (Left) or in the presence of ADAM 10 (Right) for 6 hr at 37ଌ, followed by HPLC analysis. The asterisks indicate the peptide substrates and the numbers indicate the products generated as follows: 1, EVHHQK-OH 2, LVFFAEDVGSNK-NH2 3, LVFFGEDVGSNK-NH2. (B) CD spectra of APP peptides. Measurements were carried out in the presence of 0.5% SDS.
Next we examined whether the extent of cleavage depends on the conformation of the substrate as has been described for the cleavage of APP by α-secretase (3). For this purpose, Ala-21 in Aβ(11) was replaced by Gly. This position corresponds to a naturally occurring Ala → Gly mutation at position 692 of APP770 (31), which was identified in patients with cerebral hemorrhages due to amyloid angiopathy. In vivo studies with APP770 (A692G) showed that this mutation C-terminal to the α-secretase cleavage site led to relatively more Aβ compared with p3 by partial inhibition of α-secretase (32) and to an increased alternative cleavage of the Aβ domain, probably caused by aberrant substrate recognition of α-secretase (33).
CD measurements for the octadecapeptide amide Aβ(11) in 0.5% SDS showed a spectrum characteristic for α-helical conformation, whereas a random coil conformation was observed for the peptide with the Ala → Gly mutation in position 21 of Aβ(11) (Fig. (Fig.1 1 B). The latter peptide was cleaved by ADAM 10 less efficiently than the wild-type peptide: after 30 min, only 14% of the peptide was cleaved between Lys-16 and Leu-17 compared with 28% of Aβ(11) (Table (Table1). 1 ). ADAM 10 did not cleave peptide substrates containing a cleavage site for ectodomain shedding of angiotensin-converting enzyme (34) or IL-6 receptor (35), but it cleaved a peptide substrate derived from pro-TNF-α, as reported (36, 37).
Several metalloproteinase inhibitors were examined for their ability to inhibit cleavage of Aβ(11) by purified ADAM 10. The hydroxamic acid-based inhibitor BB-3103 (38) at 100 μM completely inhibited the cleaving activity. ADAM 10 was also completely inhibited by 1 mM DTT, which suggests the existence of thiol or disulfide bonds important for α-secretase activity, and by 1 mM of the chelating agent 1,10-phenanthrolin.
Processing and Localization of ADAM 10 in HEK Cells.
To study the effect of ADAM 10 on APP cleavage in a cellular system, we used HEK cells. The ADAM 10 cDNA was cloned from a bovine kidney cDNA library, tagged at its 3′ end with a DNA sequence coding for the HA antigen, and cloned into the expression vector pcDNA3. After transfection of HEK cells and HEK cells stably expressing APP695 (HEK APP695) (29) with ADAM 10, clones were analyzed for expression and processing of the metalloprotease. For this purpose, cell lysates were immunoprecipitated with anti-HA antibody (16B12) and analyzed by SDS/PAGE and Western blot. Immunostaining revealed two immunoreactive species with apparent molecular masses of 90 kDa and 64 kDa (Fig. (Fig.2 2 A, lanes 2 and 5). The 64-kDa form is derived from the 90-kDa form by removal of the prodomain (194 aa), probably by a furin-like pro-protein convertase, which cleaves ADAMs at the sequence motif RXKR in a late Golgi compartment (39). The calculated molecular mass of the protein core of ADAM 10 after cleavage of the prodomain is 61 kDa. To investigate whether the difference between apparent and calculated molecular masses is caused by glycosylation on the putative N-glycosylation sites of ADAM 10 (24), the glycoproteins of the cell clone with the highest expression level of ADAM 10 (HEK ADAM 10) were treated with N-glycosidase F. This treatment resulted in a reduction of the molecular masses to the expected values of about 86 kDa for the precursor protein and about 61 kDa for the protein core of ADAM 10 lacking the prodomain (Fig. (Fig.2 2 A, lane 3).
Expression and deglycosylation of ADAM 10 protease (A). After transfection of HEK and HEK APP695 cells with ADAM 10 cDNA, cell lysates were immunoprecipitated with anti-HA antibody, subjected to deglycosylation with N-glycosidase F (PNGase F) (lane 3), or directly analyzed by 4% NuPAGE gel system and Western blot. (B) HEK and HEK ADAM 10 cells were surface-biotinylated. After immunoprecipitation with anti-HA antibody (16B12) and elution of the immune complexes, four-fifths of the eluate were incubated with immobilized streptavidin (lanes a), and the remaining one-fifth was directly applied to a 10% SDS/PAGE gel (lanes b) and then blotted onto PVDF membranes.
The localization of ADAM 10 in transfected HEK cells was investigated by cell surface biotinylation and confocal microscopy. For biotinylation, cells were cooled to 4ଌ and incubated for 30 min with sulfo-N-hydroxysuccinimidobiotin, a membrane-impermeant biotinylation reagent. Immunoprecipitation was performed with the anti-HA mAb. Four-fifths of collected immunoprecipitates were incubated with immobilized streptavidin to recover protease molecules localized on the plasma membrane. Fig. Fig.2 2 B shows the appearance of both the processed -kDa form of ADAM 10, as well as its 90-kDa precursor at the cell surface (lanes a). One-fifth of the immunoprecipitate that was not incubated with streptavidin predominantly showed the nonprocessed 90-kDa form and only a faint band of proteolytically activated ADAM 10 (lanes b). These results demonstrate that the proteolytically activated form of ADAM 10 is localized mainly on the cell surface, where it is able to cleave APP. The majority of ADAM 10 was found intracellularly as proenzyme. Analysis of the localization of ADAM 10 by confocal microscopy of permeabilized HEK ADAM 10 cells showed that its staining pattern largely overlapped with that of the Golgi marker 58K (Fig. (Fig.3). 3 ).
Colocalization of ADAM 10 and of a Golgi marker in HEK cells. Stably expressed ADAM 10 and the Golgi network were simultaneously immunostained. (Left) Anti-Golgi 58K protein. (Right) Anti-HA-epitope staining. The high degree of overlap of both immunoreactivities in the perinuclear region reveals a strong colocalization of ADAM 10 and the trans-Golgi marker. The scale bar is in μm.
Effect of ADAM 10 on α Secretase Cleavage of APP in HEK Cells.
The α-secretase activity in HEK ADAM 10 cells was compared with the activity found in untransfected HEK cells. The release of APPs into the medium from cells was monitored with two site-specific antibodies, 1736 and 6E10, both of which recognize the N-terminal sequence of Aβ and thus only detect APPsα, and not APPsβ. Immunoblot experiments and immunoprecipitation after metabolic labeling with 35 S were performed to determine α-secretase activity (Figs. (Figs.4 4 and and5). 5 ). Quantitative analysis of immunoblot experiments with mAb 6E10 showed that HEK cells stably expressing a high level of ADAM 10 release approximately four times more APPsα into the medium compared with untransfected HEK cells (Fig. (Fig.4, 4 , lanes 1 and 4). Increased α-secretase activity was also observed in the HEK APP695 cell clone stably expressing ADAM 10. The relative increase of APPs751α derived from endogenous APP and of APPs695α was only 2-fold (Fig. (Fig.4 4 A, lanes 7 and 8), because of the lower expression level of ADAM 10 (Fig. (Fig.2, 2 , lane 5) and the higher substrate to enzyme ratio.
Secretion of APPsα from HEK and HEK ADAM 10 cells. (A) Cells were incubated in the presence of indicated compound. After 4 hr, the medium was collected, and proteins were precipitated and subjected to immunoblot analysis with antibody 6E10 followed by a 35 S-labeled anti-mouse IgG antibody. (B) Quantitative analysis of secreted APPsα. The radioactive bands corresponding to APPsα were quantified with the Bio-Imaging analyzer model BAS-1800. The results are expressed as percentage of secreted APPsα in control HEK cells and are the averages ±SD of at least three experiments.
Effect of ADAM 10 on the production of APPsα and on the 10-kDa C-terminal fragment. (A) Cells were metabolically labeled with l -[ 35 S]methionine and [ 35 Sלysteine (200 㯌i/ml) for 5 hr. Cell media were immunoprecipitated with antibody 1736 and analyzed by SDS/PAGE in 10% gels. (B) Cell lysates were immunoprecipitated with antibody C7. The samples were separated by 10% Tris/Tricine gel (Novex) and analyzed as described. (C) Quantitative analysis of holo-APP and p10. The values of p10 were normalized to the levels of holo-APP. The results are the averages ±SD of four experiments. Statistical significance between control cells and HEK ADAM 10 cells was determined by Student’s unpaired t test (∗, P < 0.005).
Hydroxamic acid-based zinc metalloprotease inhibitors have recently been shown to inhibit the ectodomain shedding of several membrane proteins including the α-secretase cleavage of APP. No effect on the release of APPsβ has been found (40). We therefore investigated the effect of the hydroxamic acid-based inhibitor BB-3103 on HEK cells and on HEK cells overexpressing ADAM 10. BB-3103 inhibited the release of APPsα in HEK cells by about 40% (Fig. (Fig.4, 4 , lanes 1 and 3) and in HEK ADAM 10 cells by about 70% (Fig. (Fig.4, 4 , lanes 4 and 6).
The stimulation of protein kinase C by phorbol esters strongly enhances the release of APPsα and inhibits the secretion of Aβ (41, 42). To examine the effect of protein kinase C stimulation on ADAM 10 activity, HEK and HEK ADAM 10 cells were treated with 1 μM phorbol 12-myristrate 13-acetate (PMA). Immunoblot experiments showed that PMA increased APPsα release from HEK cells about 6.0-fold (Fig. (Fig.4, 4 , lanes 1 and 2). The augmented α-secretase activity from HEK ADAM 10 cells was further increased 2.5-fold (Fig. (Fig.4, 4 , lanes 4 and 5).
Similar results were observed in immunoprecipitation experiments with antibody 1736, thus the expression of ADAM 10 in HEK cells significantly increased APPsα secretion into the medium (Fig. (Fig.5 5 A). To determine whether ADAM 10 expression also increases the level of p10, cell lysates from HEK and HEK ADAM 10 cells after metabolic labeling were immunoprecipitated with antibody C7 recognizing the C-terminal part of APP. As shown in Fig. Fig.5 5 B and C, the expression of full-length APP (holo-APP) is not changed, whereas the signal for p10 is significantly stronger in HEK ADAM 10 cells than in control cells, further indicating that ADAM 10 has α-secretase activity.
Effect of a Dominant Negative ADAM 10 Mutant on α Secretase Activity.
To determine whether the human homologue of ADAM 10 is expressed in HEK cells and, therefore, might be responsible for the endogenous α-secretase activity, we performed reverse transcriptase–PCR analysis using total RNA obtained from these cells and oligonucleotides specific for ADAM 10. As shown in Fig. Fig.6 6 A, HEK cells express detectable amounts of mRNA encoding for human ADAM 10, which shows 97% identity of amino acids with bovine ADAM 10. In a control experiment, bovine ADAM 10 mRNA was detected in total RNA of MDBK cells known to express ADAM 10 (24).
Inhibition effect of a dominant negative ADAM 10 protease form (DN). (A) Endogenous expression of ADAM 10 in HEK and MDBK cells detected by reverse transcriptase–PCR. In each case a 541-bp DNA fragment could be amplified. As control, RNA from HEK cells was treated with RNase A before reverse transcription (St, DNA molecular weight marker). (B) Quantitative analysis of secreted APPsα after immunoblot analysis. The results are expressed as percentage of secreted APPsα in control HEK cells and are the averages ±SD of at least three experiments. Statistical significance between control cells and HEK˽N cells treated or untreated with PMA was determined by Student’s unpaired t test (∗, P < 0.005 ∗∗, P < 0.001).
To inhibit the endogenous α-secretase activity in HEK cells, the point mutation E384A was introduced into the zinc binding site of the bovine ADAM 10 protease. A mutant KUZ protein with a mutation at the same site acted as a dominant negative form in D. melanogaster (21). HEK cells stably expressing the mutant ADAM 10 E384A (HEK˽N) showed a substantially decreased secretion of APPs. The inhibitory effect was most apparent in PMA-treated cells: only about 25% of enzymatic activity was observed (Fig. (Fig.6 6 B). Thus, this point mutation resulted in a dominant negative form of the protease and significantly decreased constitutive and stimulated α-secretase activity.
Mechanistic and structural implications
We have successfully demonstrated that controllable cleavages of recombinant proteins can be achieved using the engineered Ssp DnaB S1 split-intein. One part of the split-intein is embedded in the precursor protein, and the complementary part of the split-intein is added in trans when needed to activate the desired cleavage reaction. In the C-cleavage design, the 144-aa IC sequence in the precursor protein could not undergo C-cleavage on its own, which demonstrates for the first time that the 11-aa IN sequence from the N-terminus of the intein is required for C-cleavage (through Asn cyclization) at the C-terminus of the intein. The IN sequence is not known to participate directly in catalyzing Asn cyclization therefore, its effect is more likely structural. In the intein crystal structure, IN is located together with the C-terminal part (including the Asn residue) of the intein in a catalytic pocket 20 therefore, the deletion of IN might have left a structural void in the catalytic pocket and thus affected cyclization of Asn. The IC was activated to undergo C-cleavage when the missing IN was added as a synthetic peptide, indicating that the IN can associate with the IC in trans to reconstitute an active intein. The IN peptide lacked an N-extein therefore, the C-cleavage reaction (Asn cyclization) must have occurred without the first two steps (acyl shift and trans-esterification) of the protein splicing mechanism.
In the N-cleavage design, the 11-aa IN sequence was embedded in the precursor protein, and the separately produced IC protein must recognize and associate with the short IN sequence in trans, in order for N-cleavage to occur. The reconstituted intein (IC plus IN) must have catalyzed an N-S acyl shift at the N-terminus of IN, and the resulting ester bond was hydrolyzed to complete the N-cleavage reaction. DTT increased the rate constant and the efficiency of the N-cleavage reaction by ∼10-fold and more than 4-fold, respectively, which is likely due to DTT promoting thiolysis of the ester bond, which occurs more rapidly than cleavage by hydrolysis.
The success of our N-cleavage design also for the first time reveals an interesting structural flexibility of the IC protein, as the IC protein could functionally assemble with the 11-aa IN sequence even when the IN was sandwiched between two large protein domains. The crystal structure of the Ssp DnaB mini-intein, from which the S1 split-intein was derived, is shaped like a disk or closed-horseshoe, 20 and the 11-aa IN sequence runs almost perpendicularly through the center of the disk-like structure [Fig. 8(A)]. The 11-aa IN sequence forms two small β-strands named β1 and β2. The β2 part of IN interacts with the β3 part of the IC protein to form an antiparallel β-sheet, which may contribute to the association between IN within the precursor protein and the IC protein. The β1 part of IN is buried deeply inside the intein structure and is enclosed by three β-strands (β5, β6, and β10) of the IC protein. It appears spatially impossible for the IN sequence to simply insert or thread itself through the central cavity of the IC protein, because in the precursor proteins tested, the IN sequence was sandwiched between two relatively large globular protein domains (the 42-kDa maltose-binding protein at the N-terminus, and proteins ranging from the 12.5-kDa thioredoxin to the 116-kDa β-galactosidase at the C-terminus). A more likely scenario is that the IC protein is structurally flexible enough to open up like a clamp so that it could saddle onto the IN sequence within the precursor protein, and then close around the IN sequence to form the active intein [Fig. 8(B)]. It is also possible that the IC protein preexists in an open-clamp structure and can change to the closed-disk-like structure only upon association with the IN sequence. The structural flexibility of the intein could be manifested by the two long β-strands (β5 and β10), which together form the backbone of the disk-like structure of the intein.
Structural modeling of IC association with IN for the N-cleavage. (A) Schematic representation of the precursor protein before and after association with the IC protein. Ribbon structures of IN (red) and IC (yellow) are adapted from a crystal structure of the Ssp DnaB mini-intein that was split into the IN and IC parts, with some of the 12 β-strands numbered according to the original crystal structure. 20 The precursor protein consists of the 11-aa IN (β-strands 1 and 2, red) sandwiched between a maltose-binding protein (MBP) and a thioredoxin (Trx). The 42-kDa MBP and the 12.5-kDa Trx are shown as round balls but not drawn to scale relative to the 17-kDa intein (IN plus IC). (B) A hypothetical model for association of IC with IN. In the first step, a transient conformational change of the IC protein loosens its structure into an open-horseshoe shape. In the second step, the IC protein saddles onto IN and closes up to form the disk-like structure of an active intein.
Our ability to precisely control the N-cleavage and the C-cleavage reactions permitted easy analysis of the reaction kinetics. Interestingly, the N-cleavage rate constant of this Ssp DnaB S1 split-intein is significantly lower than the previously reported N-cleavage rate constants of (1.0 ± 0.5) × 10 −3 s −1 for the Ssp DnaE S0 split-intein 21 and 1.9 × 10 −3 s −1 for the Sce VMA S0 split-intein. 22 This may be due to the very different amino acid sequences of these inteins, although the crystal structures of inteins are generally highly conserved. The different rate constants may be more likely due to the different split sites of these inteins, because the Ssp DnaB S1 split-intein is split at the S1 site near the N-terminus of the intein sequence, whereas the Ssp DnaE split-intein and the Sce VMA split-intein are split at the S0 site closer to the middle of the intein sequence. Different split sites have been noted to affect the rate constant of trans-splicing reactions. The Ssp DnaB S0 split-intein and the Ssp DnaB S1 split-intein, which were derived from the same natural intein, showed trans-splicing rate constants of (9.9 ± 0.8) × 10 −4 s −1 and (4.1 ± 0.2) × 10 −5 s −1 , respectively, 19 , 22 which differed by more than an order of magnitude. Unlike the N-cleavage rate constant discussed above, the C-cleavage rate constants of this Ssp DnaB S1 split-intein are quite comparable to the previously reported C-cleavage rate constants of (1.9 ± 0.9) × 10 −4 s −1 for the natural Ssp DnaE split-intein and (1.0–1.7) × 10 −3 s −1 for the synthetic Sce VMA split-intein, 21-23 although these split-inteins have very different amino acid sequences and very different split sites. We found that the C-cleavage rate constant could be increased by ∼10-fold by substituting the Cys1 residue to the less nucleophilic Ser residue. Although the residues at the N-terminal splice junction are not known to directly participate in Asn cyclization, it appears that even small changes in amino acid side chains at the intein N-terminus can substantially affect rearrangement of chemical bonds at the C-terminal splice junction.
Potential uses and advantages of the controllable cleavages
Advantages of intein-based protein cleavage methods, compared to others such as protease-based methods, have been noted previously, 11 and our methods using intein fragments retain many of these advantages. For example, the N-cleavage method may be used to generate an activated thioester at the C-terminus of a target protein so that the target protein can be joined with another protein or peptide having an N-terminal Cys residue, using the EPL method. 8-10 Unlike protease-based methods that cleave on the C-terminal side of specific recognition sequences, our intein-based N-cleavage method cleaves on the N-terminal side of the recognition sequence (IN) and thus allows removal of the affinity purification domain (or tag) placed on the C-terminus of the target protein. The intein-based N- and C-cleavage methods may also be used together on a single target protein to produce precise and tag-free ends at both the N- and the C-termini, or to achieve cyclization of the target protein (ligation of the N- and C-termini) using the EPL approach. We have demonstrated that the C-cleavage method can be used to generate an N-terminal Cys residue on a target protein, which is needed for EPL, although the Ssp DnaB intein is naturally followed by a Ser residue. Another advantage of these intein-based methods, unlike some protease-based methods, is that inteins are believed to pose no risk of nonspecific cleavage at unintended locations.
Compared to existing intein-based methods that use contiguous inteins, our methods using the S1 split-intein completely avoid any spontaneous cleavage during expression and purification of the precursor protein. This is a significant advantage, because spontaneous cleavages often result in lower yields of the purified protein. In previous reports using contiguous inteins, unwanted spontaneous cleavages have been observed in vivo at various levels and could be as high as 90%. 9 In our C-cleavage method, the 11-aa IN peptide may not be overly expensive and laborious to use, due to the small size of the peptide. For example, cleavage of 100 mg of a 50 kDa-precursor protein may require as little as 20 mg of the IN peptide, if the peptide is used at ∼10-fold molar excess over the precursor protein to drive the cleavage reaction. The extra cost of 20 mg IN may easily be compensated by an increase in protein yield due to the prevention of spontaneous cleavage observed with other intein-based methods. This cost-effectiveness may be particularly true when producing high value proteins for research or pharmaceutical uses, when using more expensive producing cells (e.g. mammalian tissue culture), or when spontaneous cleavages are not tolerated. Lastly, the small IN peptide can be stably stored and easily removed from the cleaved proteins through simple dialysis.
The recombinant precursor proteins in this study had either a maltose-binding protein or a chitin-binding domain as the affinity binder for easy purification, but potentially other affinity binders such as the His-tag and the GST-tag may be used. We showed that the C-cleavage can also occur when the precursor protein is bound to chitin beads and incubated with the IN peptide therefore, this C-cleavage method may be used in a single-step purification of recombinant proteins, using a process similar to that of the IMPACT method. 5 In such a process, cell lysate containing the precursor protein is passed through an affinity column, unbound proteins are washed away, the IN peptide is added to the column to activate the C-cleavage, and the protein of interest is released (cleaved) from the column in a pure form. We have shown the feasibility of this purification strategy by preparing two recombinant proteins (thioredoxin and eGFP) in good yields (6 and 3 mg/L of culture). Our demonstration of peptide-triggered C-cleavage with various different target proteins in solution further shows that the purification approach could be a valuable tool for the preparation of recombinant proteins.
Both N- and C-cleavage reactions presented here reached efficiencies of >90% under optimal conditions, which is comparable to or higher than the cleavage efficiencies of previous methods using contiguous inteins. The N-cleavage method in this study also showed a much higher (∼95%) cleavage efficiency compared to the previously reported IMPACT method using contiguous inteins, 5 probably because the shorter IN sequence embedded in the precursor protein less likely causes protein misfolding.
Background:Developments in ‘soft’ ionisation techniques have revolutionized mass-spectro-metric approaches for the analysis of protein structure. For more than a decade, such techniques have been used, in conjuction with digestion b specific proteases, to produce accurate peptide molecular weight ‘fingerprints’ of proteins. These fingerprints have commonly been used to screen known proteins, in order to detect errors of translation, to characterize post-translational modifications and to assign diulphide bonds. However, the extent to which peptide-mass information can be used alone to identify unknown sample proteins, independent of other analytical methods such as protein sequence analysis, has remained largely unexplored.
Results: We report here on the development of the molecular weight search (MOWSE) peptide-mass database at the SERC Daresbury Laboratory. Practical experience has shown that sample proteins can be uniquely identified from a few as three or four experimentally determined peptide masses when these are screened against a fragment database that is derived from over 50 000 proteins. Experimental errors of a few Daltons are tolerated by the scoring algorithms, thus permitting the use of inexpensive time-of-flight mass spectrometers. As with other types of physical data, such as amino-acid composition or linear sequence, peptide masses provide a set of determinants that are sufficiently discriminating to identify or match unknown sample proteins.
Conclusion: Peptide-mass fingerprints can prove as discriminating as linear peptide sequences, but can be obtained in a fraction of the time using less protein. In many cases, this allows for a rapid identification of a sample protein before committing it to protein sequence analysis. Fragment masses also provide information, at the protein level, that is complementary to the information provided by large-scale DNA sequencing or mapping projects.
MATERIALS AND METHODS
All of the mutant strains of Chlamydomonas reinhardtii used in this study have been described previously (see Table 2). The oda2 allele used was pf28 (Mitchell and Rosenbaum, 1985). Cells were maintained on minimal medium using standard procedures (Harris, 1989).
Cell Cytoplasmic Extract
Chlamydomonas cells were grown in 500 ml of liquid M medium (Sager and Granick, 1953) with aeration in continuous light to a density of 10 6 cells/ml, harvested by centrifugation (550 × g for 6 min at 22°C), and resuspended in ice-cold HMDEK (10 mM HEPES, 5 mM MgSO4, 1 mM DTT, 0.1 mM EDTA, 25 mM potassium chloride, pH 7.4) to a total of 500 μl. The suspension was transferred to a 1.5-ml microfuge tube that contained an equal volume of acid-washed glass beads (1 mm) and vortexed at setting 6.5 on a Genie II vortexer for 1 min. Cell suspensions were then centrifuged using a Beckman L8 centrifuge at 48,000 ×g, at 4°C for 2 h. Supernatants were either used for immunoprecipitation as described below or mixed with 0.25 volume of 4× sample buffer (8% SDS, 40% glycerol, 125 mM Tris-HCl, pH 6.8, with Pyronin Y added as tracking dye) and β-mercaptoethanol, to a final concentration of 0.7 M, and stored at −20°C for SDS-PAGE. Pellets were resuspended in 500 μl HMDEK, mixed with 0.25 volume 4× sample buffer, and stored at −20°C for SDS-PAGE. Protein concentration was determined by the Bradford dye binding method using BSA as a standard (Bradford, 1976).
Axonemes were isolated by the method of Witman et al.(1978). Cells were grown in 500 ml of liquid M medium (Sager and Granick, 1953) with aeration in continuous light to a density of 10 6 cells/ml, harvested by centrifugation at 550 ×g for 6 min at 22°C, washed with 10 mM HEPES, pH 7.4, centrifuged again, and resuspended in 10 ml HMDS (10 mM HEPES, pH 7.4, 5 mM MgSO4, 1 mM DTT, and 4% sucrose). Resuspended cells were deflagellated with 400 μl 50 mM dibucaine (CIBA Pharmaceutical, CIBA-GEIGY, Summit, NJ) and diluted with 10 ml ice-cold HMDS containing 2 mM EGTA and 2 mM phenylmethylsulfonyl fluoride, and cell bodies were removed by centrifugation at 4°C for 7 min at 1,550 ×g. The supernatant was collected and recentrifuged as above. Cell-free supernatant was then centrifuged at 31,000 ×g to pellet axonemes, which were resuspended in HMDEK and an equal volume of 2× sample buffer. β-Mercaptoethanol was added to a final concentration of 0.7 M, and samples were stored at −20°C.
SDS-PAGE and Western Blotting
Samples were prepared and run with Tris-glycine-buffer (Laemmli, 1970) in 5% stacking gels and 5, 7, or 12% separating gels (designated in text) prepared from stocks that contained 30% acrylamide and 0.4% bis-acrylamide. Broad Range protein standards (New England Biolabs, Beverly, MA) of 212, 158, 116, 97.2, 66.4, 55.6, and 42.7 kDa were used, and gels were either stained with Coomassie Blue to show total protein or transferred to immobilon membrane (Millipore, Bedford, MA) for Western blotting following the recommendations ofBurnette (1981). Gels were soaked in transfer buffer (25 mM Tris, 192 mM glycine, 10% methanol) for 10 min and transferred either at 200 mA for 12 h (Figures 2, 3, 4A, and 5) or at 300 mA for 6 h (Figures 4B and 6). Protein standard lanes were separated from sample lanes and stained with amido black. Antibody binding and detection were performed as directed in the POD chemiluminescence kit (Boehringer Mannheim, Indianapolis, IN). Briefly, transferred blots were blocked with 1% POD blocking solution for 1 h at room temperature, incubated with the primary antibody in 0.5% POD blocking solution for 3 h at room temperature, washed with TBST (50 mM Tris base, 150 mM NaCl, pH 7.5, with 0.1% Tween-20 [vol/vol]) 2 × 10 min, washed with 0.5% POD blocking solution 2 × 10 min, incubated with secondary antibody in 0.5% POD blocking solution for 1 h at room temperature, washed with TBST four times for 10 min, and then incubated with developing solution for 1 min and exposed to Biomax film (Eastman Kodak, Rochester, NY). Antigen quantitation was estimated by comparison with a blot of an antigen dilution series (2, 1, 0.8, 0.6, 0.4, 0.2, 0.1 x WT control) processed in parallel.
Fig. 2. Specificity of antibody B3B. WT andoda11 flagella, run on 5% gels and blotted to PVDF membrane, were probed with C11.6 (left panel), which detected equal levels of HCβ in both samples. A parallel blot probed with antibody B3B (right panel) detected a single band in WT flagella that is missing in flagella of HCα assembly mutant oda11. Antibodies were detected with a peroxidase-linked secondary antibody and chemiluminescent detection.
Anti-HCβ mAb C11.6 was concentrated by ammonium sulfate precipitation from hybridoma culture supernatants and used at a 1:100 dilution. It was generated from the same fusion as C11.13 (Mitchell and Rosenbaum, 1986). Anti-IC70 mAb 1869A ascites was used at 1:2000 dilution. Anti-IC78 mAb 1878A hybridoma culture supernatant was kindly donated by Dr. G. Witman (King et al., 1985) and was used at a 1:2 dilution. Anti-HCα polyclonal antibody B3B was produced in a rabbit by immunization with a purified bacterial fusion protein. A 1-kilobase (kb) PmlI/HpaI fragment of HCα cDNA pBcA6 (Mitchell and Brown, 1997), which encodes amino acids 512–838 of HCα (a region unrelated to other DHC sequence and that contains the HCα EPAA repeat element), was cloned into vector pGEX-4T-2 (Pharmacia Biotech, Piscataway, NJ) at a SmaI site and was transformed into DH5αF′ E. coli. Fusion protein expression was induced for 3 h with 0.1 mM isopropyl-β-thiogalactopyranoside at 37°C. Fusion protein was solubilized and purified by the method ofFrangioni and Neel (1993), run on an SDS-PAGE gel, transferred to nitrocellulose, and visualized with Ponceau S. Nitrocellulose strips containing the protein of interest were dissolved in dimethyl sulfoxide, mixed with adjuvant, and used for immunization. Specific antibodies were affinity purified from whole sera using Western blots of the fusion protein (Frangioni and Neel, 1993) and were used at a dilution of 1:50. Anti-HCγ mAb 25–8, kindly donated by Dr. G. Piperno (King and Witman, 1988), was used at a 1:10 dilution. Peroxidase-labeled goat anti-mouse or goat anti-rabbit secondary antibodies (Bio-Rad Laboratories, Hercules, CA) were used at a 1:6000 dilution.
Cell extracts (0.5 ml) were mixed in a 15-ml conical tube with an equal volume of ice-cold immunoprecipitation (IP) buffer (HMDEK, 75 mM NaCl, 0.01% thimersol, 0.5 mM PMSF, 3% BSA, 0.1% Triton X-100, pH 7.5) and preabsorbed with 25 μl of 50/50 (vol/vol) protein A agarose in IP buffer for 30 min on ice. mAb anti-HCβ antibody C11.6 was added to preabsorbed extracts and incubated for 3–4 h at 4°C. A 100-μl volume of 50/50 (vol/vol) protein A agarose (Sigma Chemical, St. Louis, MO) in IP buffer was added, and the tubes were mixed gently for 1 h. Agarose beads were washed three times with IP buffer containing 0.05% Triton X-100. Immune complexes were eluted by addition of 2× sample buffer containing 0.7 M β-mercaptoethanol and incubation for 2 min in boiling water. The quantity of HCβ immunoprecipitated with antibody C11.6 from wild-type andoda mutant cytoplasmic extracts was determined from preliminary Western blots. Subsequent loads of immunoprecipitate samples were adjusted to include equal amounts of HCβ.
Partial Acid Hydrolysis
Western blots of proteins subjected to partial acid hydrolysis were generated by a modification of the method described by Clevelandet al. (1977). Briefly, cytoplasmic extracts or whole axonemes were run on an SDS-PAGE gel along with molecular weight markers. The gel was stained in 0.1% Coomassie blue, 50% methanol, and 10% acetic acid for 30 min and destained in 5% methanol and 10% acetic acid for 45 min. Bands of ∼70 kDa molecular mass were cut from the gel and soaked 30 min in 0.125 M Tris/HCl, pH 6.8, 0.1% SDS, 1 mM EDTA. Slices were then lyophilized and either stored frozen at −20°C (controls) or incubated with 70% formic acid for 16 h at 37°C, washed with 50% methanol, and lyophilized. Gel slices were then rehydrated with buffer (1% SDS, 10 mM Tris/HCl, pH 8, 0.1% β-mercaptoethanol, and 10% glycerol) for 6 h, pushed into the bottom of an SDS-PAGE well, and overlayed with 10 μl of Tris/HCl buffer containing 20% glycerol for electrophoresis and transfer to PVDF membranes.
9.9 Test Prep for AP Courses
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This text is based on Openstax Biology for AP Courses, Senior Contributing Authors Julianne Zedalis, The Bishop's School in La Jolla, CA, John Eggebrecht, Cornell University Contributing Authors Yael Avissar, Rhode Island College, Jung Choi, Georgia Institute of Technology, Jean DeSaix, University of North Carolina at Chapel Hill, Vladimir Jurukovski, Suffolk County Community College, Connie Rye, East Mississippi Community College, Robert Wise, University of Wisconsin, Oshkosh
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