s2fieldsofstudy:
Biology
Computer Science
Chemistry

text:
Definition of the p53 functional domains necessary for inducing apoptosis.

The p53 protein contains several functional domains necessary for inducing cell cycle arrest and apoptosis. The C-terminal basic domain within residues 364-393 and the proline-rich domain within residues 64-91 are required for apoptotic activity. In addition, activation domain 2 within residues 43-63 is necessary for apoptotic activity when the N-terminal activation domain 1 within residues 1-42 is deleted (DeltaAD1) or mutated (AD1(-)). Here we have discovered that an activation domain 2 mutation at residues 53-54 (AD2(-)) abrogates the apoptotic activity but has no significant effect on cell cycle arrest. We have also found that p53-(DeltaAD2), which lacks activation domain 2, is inert in inducing apoptosis. p53-(AD2(-)DeltaBD), which is defective in activation domain 2 and lacks the C-terminal basic domain, p53-(DeltaAD2DeltaBD), which lacks both activation domain 2 and the C-terminal basic domain, and p53-(DeltaPRDDeltaBD), which lacks both the proline-rich domain and the C-terminal basic domain, are also inert in inducing apoptosis. All four mutants are still capable of inducing cell cycle arrest, albeit to a lesser extent than wild-type p53. Interestingly, we have found that deletion of the N-terminal activation domain 1 alleviates the requirement of the C-terminal basic domain for apoptotic activity. Thus, we have generated a small but potent p53-(DeltaAD1DeltaBD) molecule. Furthermore, we have determined that at least two of the three domains (activation domain 1, activation domain 2, and the proline-rich domain), are required for inducing cell cycle arrest. Taken together, our results suggest that activation domain 2 and the proline-rich domain form an activation domain for inducing pro-apoptotic genes or inhibiting anti-apoptotic genes. The C-terminal basic domain is required for maintaining this activation domain competent for transactivation or transrepression.

p53 is frequently mutated in cancers. Mutations in the p53 DNA binding domain or certain mutations in the nuclear localization signal and tetramerization domain that indirectly affect DNA binding abrogate or diminish p53 activity in cell cycle arrest and apoptosis (1,5). The proline-rich domain has been shown to be required for efficient growth suppression (12). Recent experiments indicate that the proline-rich domain is necessary for apoptosis but not cell cycle arrest (16 -18). In addition, the proline-rich domain plays an important role in the induction of several endogenous target genes, but is not required for activation of the exogenously introduced promoters of these target genes (17). These results suggest that the proline-rich domain may participate in the induction of cellular target gene(s) responsible for mediating apoptosis. However, the role of other p53 functional domains (especially the Nterminal activation domain 1 and the C-terminal basic domain) in apoptosis is still not certain. Earlier reports have shown that in some experimental protocols (19 -21) including our own (22), p53 transactivation activity is dispensable for apoptosis. It should be noted that this conclusion is based at least in part on the observation that an activation domain 1-deficient mutant (a double point mutation at residues 22-23, AD1 Ϫ ) 1 is capable of inducing apoptosis (21,22). Recently, we and others have shown that p53-(AD1 Ϫ ) contains an intact activation domain 2 (9 -11), and therefore, p53-(AD1 Ϫ ) is still competent in transactivation (10). Furthermore, when both activation domain 1 and activation domain 2 are mutated (a quadruple point mutation at residues 22-23 and 53-54, AD1 Ϫ AD2 Ϫ ), the resulting protein is inert in transactivation and in inducing cell cycle arrest and apoptosis (9 -11).
The C-terminal basic domain has been subjected to extensive analysis, and all evidence suggests that the basic domain is a regulatory domain. This basic domain can regulate the DNA binding activity when it is phosphorylated (1,5), acetylated (23)(24)(25), deleted (26), or associated with anti-p53 antibody (26,27) or peptides derived from the C terminus of p53 (28,29). Interestingly, the mechanism by which these latter peptides enhance p53 DNA binding activity is the ability of the peptides to interact with three separate domains in p53, that is, the proline-rich domain (30), the DNA binding domain (31), and the C-terminal basic domain (30,31). The C-terminal basic domain also interacts with several cellular proteins, such as TFIIH subunits XPB and XPD (32,33), and Werner syndrome protein (WRN) (34,35), which all lead to efficient induction of p53-mediated apoptosis. These results support a hypothesis that the C-terminal basic domain is a negative regulatory do-main whose effect on the DNA binding activity can be alleviated by interacting with other cellular proteins, peptides derived from the p53 C terminus, or other modifications. However, several groups have shown that p53-(⌬BD), which lacks the C-terminal basic domain, has a reduced ability to induce several cellular target genes and becomes incapable of inducing apoptosis (22,32,36). These results suggest that the C-terminal basic domain can regulate p53 activity both positively and negatively.
In this study, we show that activation domain 2 and the proline-rich domain form an activation domain for inducing pro-apoptotic genes or inhibiting anti-apoptotic genes. The Cterminal basic domain is required for maintaining this activation domain competent for transactivation or transrepression. We also found that an activation domain capable of inducing at least partial cell cycle arrest can be formed by activation domain 1 plus activation domain 2, activation domain 1 plus the proline-rich domain, or activation domain 2 plus the prolinerich domain. The ability of these activation domains to induce cell cycle arrest can be enhanced by the presence of the Cterminal basic domain.
The above mutant p53 cDNAs were cloned separately into a tetracycline-regulated expression vector, pUHD10-3, at its EcoRI site (37), and the resulting plasmids were used to generate cell lines that inducibly express p53.
Growth Rate Analysis, Trypan Blue Dye Exclusion Assay, DNA Histogram Analysis, and Annexin V Staining-Growth rate analysis, trypan blue dye exclusion assay, and DNA histogram analysis were performed as described previously (10,17,22). Propidium iodide and RNase A were purchased from Sigma. Fluorescein isothiocyanate-labeled annexin V was purchased from Roche Molecular Biochemicals, and staining was performed as described by the manufacturer.
RNA Isolation and Northern Blot Analysis-Total RNA was isolated using Trizol reagents (Life Technologies, Inc.). Northern blot analysis was performed as described (10). The p21, BAX, and glyceraldehyde-3phosphate dehydrogenase probes were prepared as described previously (10).

RESULTS
The Activity of Activation Domain 2 Is Necessary for Inducing Apoptosis-Previously, we have shown that the activity of activation domain 2 is required for inducing apoptosis when a double point mutation at residues 22-23 or deletion of the N-terminal 42 amino acid residues renders activation domain 1 dysfunctional (10). To further determine the function of activation domain 2 in apoptosis, we generated an activation domain 2-deficient mutant, p53-(AD2 Ϫ ), which contains a double point mutation at residues 53-54. We then established several cell lines that inducibly express this mutant in p53-null H1299 lung carcinoma cells. Western blots from two representative cell lines, p53-(AD2 Ϫ )-6 and -8, are shown in Fig. 1A. After normalization to the levels of actin protein expressed, we found that the levels of p53 protein in p53-(AD2 Ϫ )-6 and -8 cells were comparable with that in p53-3 and HA⅐p53-15 cells, which express wild-type p53 and HA-tagged wild-type p53, respectively (Fig. 1A, upper two panels, compare lanes 5-8 with lanes [1][2][3][4]. To determine the transcriptional activity of p53-(AD2 Ϫ ), we measured the level of p21 protein induced by p53-(AD2 Ϫ ). Surprisingly, we found that the ability of p53-(AD2 Ϫ ) to induce p21 was severely diminished (Fig. 1A, p21 panel, lanes [5][6][7][8]. These results are similar to that observed for the activation domain 1-deficient mutant (6,10,22). In contrast, p21 was strongly induced by wild-type p53 and HA-tagged wild-type p53 (Fig. 1A, p21 panel, lanes 1-4).
To determine the activity of p53-(AD2 Ϫ ) in H1299 cells, the growth rate of p53-(AD2 Ϫ )-6 cells was determined over a 5-day period. When induced to express p53-(AD2 Ϫ ), cells failed to multiply (Fig. 1B), but visible microscopic cell death was not significantly increased (data not shown).
To determine whether the growth suppression by p53-(AD2Ϫ) is because of cell cycle arrest, apoptosis, and/or both, we performed a DNA histogram analysis and an annexin V staining assay. When induced to express the mutant p53-(AD2 Ϫ ) for three days, we found that the percentage of cells in S phase decreased from 35 to 8% whereas cells in G 1 increased from 49 to 75%, suggesting that p53-(AD2 Ϫ ) arrested cells primarily in G 1 (Fig. 1, D-E). However, no apparent apoptosis was detected by either DNA histogram analysis (Fig. 1, D-E) or annexin V staining (Fig. 1, F-G). Thus, the activity in activation domain 2 is necessary for inducing apoptosis. As a positive control, we analyzed p53-3 and HA⅐p53-15 cells. When induced to express wild-type or HA-tagged p53 for three days, we found that both p53-producing cells were arrested primarily in G 1 and underwent apoptosis, consistent with previous reports (10,22). We also analyzed p53-(AD2 Ϫ ⌬BD)-9 cells. We found that no significant apoptosis was observed, and cells primarily arrested in G 1 when induced to express p53-(AD2 Ϫ ⌬BD) (data not shown).
To determine the activity of the entire activation domain 2 (residues 43-62), we generated p53-(⌬AD2), which lacks the entire activation domain 2 and p53-(⌬AD2⌬BD), which in turn lacks activation domain 2 and the C-terminal basic domain. We then established several cell lines that inducibly express p53-(⌬AD2) and p53-(⌬AD2⌬BD), respectively (Fig. 2, A and C). We found that p53-(⌬AD2) and p53-(⌬AD2⌬BD) suppressed cell proliferation (Fig. 2, B and D), albeit to a lesser extent than p53-(AD2 Ϫ ) and p53-(AD2 Ϫ ⌬BD) (Fig. 1, B and C). Furthermore, we found that cells were arrested primarily in G 1 but did not undergo apoptosis when induced to express these p53 mutants (data not shown, Table I). However, we found that p21 was not significantly induced (Fig. 2, A and C), suggesting that p53-dependent cell cycle arrest in G 1 can be mediated by a gene(s) other than p21.
The Proline-rich Domain Contributes to the Ability of p53 to Induce Cell Cycle Arrest-Previously, we and others have shown that the proline-rich domain (16 -18) and the C-terminal basic domain (22,32) are necessary for inducing apoptosis but not cell cycle arrest. To determine whether both domains are dispensable for inducing cell cycle arrest, we generated p53-(⌬PRD⌬BD), which lacks both the proline-rich domain and the C-terminal basic domain. We then established several cell lines that inducibly express this mutant. Western blots from three representative cell lines, p53-(⌬PRD⌬BD)-2, -6, and -7, are shown in Fig. 2E. We found that the level of p53 expressed in p53-(⌬PRD⌬BD)-2 cells was comparable with that in p53-3, HA⅐p53-15, and p53-(⌬BD)-1 but slightly lower than that in p53-(⌬PRD)-5, which inducibly expresses a p53 mutant lacking FIG. 1. The activity of activation domain 2 is necessary for inducing apoptosis. A, levels of p53, p21, and actin were assayed by Western blot analysis in cell lines as shown above the blots. Cell extracts were prepared from non-induced cells (Ϫ) and cells induced to express p53 for 24 h (ϩ). HA-tagged p53 was detected with 12CA5 antibody. p53 was detected with anti-p53 monoclonal antibody Pab240. p21 was detected with anti-p21 monoclonal antibody (Ab-1). Actin was detected with anti-actin polyclonal antibody. B and C, growth rates of p53-(AD2 Ϫ )-6 and p53-(AD2 Ϫ ⌬BD)-9 cells in the absence (छ) or presence (Ⅺ) of p53 over a 5-day period. D and E, DNA content was quantified by propidium iodide staining of fixed cells that were non-induced (Ϫ p53) or induced (ϩ p53) to express p53-(AD2 Ϫ ) for 3 days. F and G, apoptotic cells were quantified by propidium iodide-annexin V staining of cells that were non-induced (Ϫ p53) or induced (ϩ p53) to express p53-(AD2 Ϫ ) for 3 days.
p53 Functional Domains for Apoptosis the proline-rich domain (Fig. 2E, p53 panel). To determine whether p21 can be induced, we found that p53-(⌬PRD⌬BD) was much less potent in inducing p21 than wild-type p53, HA-tagged p53, p53-(⌬BD), or p53-(⌬PRD) (Fig. 2E, p21 panel). However, when the DNA binding activity was determined in vitro, we found that p53-(⌬PRD⌬BD) was as potent as wildtype p53 in binding to the ribosomal gene cluster p53 response element (data not shown). This suggests that deletion of both the proline-rich domain and the C-terminal basic domain does not affect the activity of the p53 DNA binding domain. Growth rate analysis showed that p53-(⌬PRD⌬BD) had a much reduced ability to suppress cell proliferation (Fig. 2F). In addition, DNA histogram analysis and annexin V staining assay showed that a partial arrest in G 1 , but no apoptosis, was detected in p53-(⌬PRD⌬BD)-2 cells (data not shown).
p53-(⌬AD1⌬BD) Is Small but Potent in Inducing Cell Cycle Arrest and Apoptosis-We and others have shown that p53-(⌬BD), which lacks the C-terminal basic domain, is inactive in inducing apoptosis (22,32,36) whereas p53-(⌬AD1), which lacks activation domain 1 (residues 1-42), is very active (10). To determine whether the C-terminal basic domain is necessary for p53-(⌬AD1) to induce apoptosis, we generated p53-(⌬AD1⌬BD), which lacks activation domain 1 and the C-terminal basic domain. We then established several cell lines that inducibly express p53-(⌬AD1⌬BD). Western blots from three representative cell lines, p53-(⌬AD1⌬BD)-3, -6, and -7, are shown in Fig. 3A. We found that the level of p53 expressed in these cells was comparable with that in p53-3, HA⅐p53-15, and p53-(⌬BD)-1 cells, but lower than that in p53-(⌬AD1)-2 cells (Fig. 3A, p53 panel). p53-(⌬AD1)-2 cells are derived from H1299 cells that inducibly express p53-(⌬AD1), which lacks activation domain 1 (10). We found that p53-(⌬AD1⌬BD) retained the ability to induce p21. Induction of p21 by p53-(⌬AD1⌬BD) was greater than induction by p53-(⌬AD1) but less than induction by wild-type p53 and p53-(⌬BD) (Fig. 3A, p21  panel). Growth rate analysis showed that cells failed to multiply when induced to express p53-(⌬AD1⌬BD) (Fig. 3, B and C). Microscopic examination showed that the p53-expressing cells detached from plates and shrank to form apoptotic bodies (data not shown). DNA histogram analysis showed that the percentage of cells in S phase decreased from 35 to 11% but the percentage of cells in G 1 increased from 55 to 75%, suggesting that these cells arrested primarily in G 1 (Fig. 3, D-E). We also found that the number of cells with a sub-G 1 DNA content was not significantly increased. However, when stained for annexin V, we found that the percentage of stained cells increased from 7 to 31%, suggesting that these cells also underwent apoptosis (Fig. 3, F-G).
Growth rate analysis showed that cells failed to multiply when induced to express wild-type p53 or p53-(⌬AD1⌬BD) (Fig. 4, B-C). Microscopic examination showed that the p53-expressing cells detached from plates and shrank to form apoptotic bodies (data not shown). DNA histogram analysis showed that the percentage of cells that had a sub-G 1 DNA content was increased from 3 to 37% by wild-type p53 (Fig. 4, D-E) and from 4 to 49% by p53-(⌬AD1⌬BD) (Fig. 4, H-I). In addition, annexin V staining assay showed that the percentage of the annexin V-stained cells was increased from 7 to 28% by wild-type p53 and from 9 to 29% by p53-(⌬AD1⌬BD). These data indicate that p53-(⌬AD1⌬BD) is a potent apoptotic inducer. At Least Two of the Three Domains, i.e. Activation Domain 1, Activation Domain 2, and the Proline-rich Domain Are Required for Inducing Cell Cycle Arrest-To further define the role of activation domain 1, activation domain 2, the prolinerich domain, and the C-terminal basic domain in inducing cell cycle arrest and apoptosis, we generated six p53 mutants that are dysfunctional in two or three of the four functional domains (Fig. 5). We then established several cell lines that inducibly express these p53 mutants individually (Fig. 5). These are p53-(⌬AD1AD2 Ϫ ⌬BD),  5E. The level of p53 expressed in some of these mutant p53producing cells was comparable with or higher than that in p53-3 cells (Fig. 5, A-E, p53 panel). However, none of these mutants were capable of inducing p21 (Fig. 5, A-E, p21 panel). In addition, cell cycle arrest and apoptosis were not detected by growth rate and DNA histogram analyses and annexin V staining assay (data not shown). These data suggest that at least two of the three domains (activation domain 1, activation domain 2, and the proline-rich domain) are required for p53 activity.
Regulation of p21 and BAX by p53 Mutants-To determine the ability of various p53 mutants that lack activation domain 1, activation domain 2, and/or the C-terminal basic domain in inducing p21 and BAX, we performed a Northern blot analysis (Fig. 6). We found that wild-type p53 was very active (lanes 1-2). p53-(R249S), a tumor-derived mutant that is defective in the DNA binding domain, was nearly inert (lanes 3-4). Although deletion of the C-terminal basic domain renders p53 constitutively active in binding to DNA in vitro (26), the ability of p53-(⌬BD) to induce p21 and BAX was approximately 2-fold less efficient than that of wild-type p53 (compare lanes 1-2 and   FIG. 3. The C-terminal basic domain is not necessary for apoptosis when activation domain 1 is absent. A, levels of p53, p21, and actin were assayed by Western blot analysis in cell lines as shown in the absence (Ϫ) or presence (ϩ) of p53 for 24 h. Antibodies used were as described in the legend to Fig. 1. B and C, growth rates of p53-(⌬AD1⌬BD)-6 and p53-(⌬AD1⌬BD)-7 cells in the absence (Ⅺ) or presence (छ) of p53 over a 5-day period. D and E, DNA content was quantified by propidium iodide staining of fixed cells that were non-induced (Ϫ p53) or induced (ϩ p53) to express p53-(⌬AD1⌬BD) for 3 days. F and G, apoptotic cells were quantified by propidium iodideannexin V staining of cells that were noninduced (Ϫ p53) or induced (ϩ p53) to express p53-(⌬AD1⌬BD) for 3 days.
p53 Functional Domains for Apoptosis [7][8]. p53-(AD1 Ϫ ) (lanes 5-6), p53-(⌬AD1) (lanes 9 -10), p53-(AD2 Ϫ ) (lanes [13][14], and p53-(AD2Ϫ⌬BD) (lanes [15][16] were extremely weak in inducing p21 and BAX (2-fold or less). It should be mentioned that p53-(AD2 Ϫ ) is extremely potent in inducing G 1 arrest (see Fig. 1, D-E), suggesting that a gene(s) other than p21 is responsible for this. Furthermore, when activation domain 1 and the basic domain were deleted, the ability of p53-(⌬AD1⌬BD) to induce p21 and BAX was partially restored (lanes [11][12], consistent with the result detected by Western blot analysis (Fig. 3A). DISCUSSION p53 induces apoptosis but the underlying mechanism remains unclear. To determine this mechanism, two major questions need to be addressed. What domains in p53 are required and is p53 transcriptional activity necessary for inducing apoptosis? Previous attempts to answer these questions have been inconclusive, because different experimental systems have been used (1,2). These include various types of cell lines and methods to express p53 (transient versus stable, ectopic versus inducible) and different types of p53 mutants (temperature-sensitive mutant versus wild-type p53; point mutations versus deletion mutations). To avoid these problems, we have applied the tetracycline inducible expression system to stably express various p53 mutants in p53-null H1299 cells. On the basis of the results obtained in this study (Table I) and several previous studies (11, 12, 16, 32, 36, 38 -40), including our own (10,17,22), we propose the following model for p53 functional domains in apoptosis (Fig. 7). First, p53 DNA binding activity is necessary for apoptosis because mutants that are defective in the DNA binding and tetramerization domains are inert. Second, activation domain 2 and the proline-rich domain can form an activation domain for transactivating pro-apoptotic genes or transrepressing anti-apoptotic genes, because mutation or deletion in either one of the domains abrogates the apoptotic activity. Third, activation domain 1 is not required because deletion of or mutation in activation domain 1 (p53-(⌬AD1), p53-(AD1 Ϫ )) has little effect on apoptosis. Fourth, the C-terminal basic domain is necessary for maintaining p53 competent in inducing apoptosis, probably by relieving the inhibitory activity of activation domain 1, because p53-(⌬AD1⌬BD), but not p53-(⌬BD), is capable of inducing apoptosis.
Several p53 inducible genes, such as BAX (41), KILLER/ DR5 (42), and several PIGs (43), may participate in the apoptotic process. These genes can be induced by either p53-(⌬PRD) (17) or p53-(AD2 Ϫ ) (data not shown), both of which are active in inducing cell cycle arrest but not apoptosis, suggesting that these genes are not required or insufficient for inducing apoptosis. Recent evidence has shown that p53 can repress specific genes, such as MAP4 (44). It is possible that transrepression of anti-apoptotic genes plays an important role in p53-mediated  H and I). F, G, J, and K, apoptotic cells were quantified by propidium iodide-annexin V staining of cells that were non-induced (Ϫ p53) or induced (ϩ p53) for 3 days to express p53 (F and G) or p53-(⌬AD1⌬BD) (J and K). p53 Functional Domains for Apoptosis apoptosis. Therefore, the cell lines that inducibly express the p53 mutants described in this study, especially p53-(⌬AD1⌬BD), can be used to identify and determine whether a cellular gene is necessary for mediating p53-dependent apoptosis.
p53 transcriptional activity has been shown to be necessary for inducing cell cycle arrest (1,2,4,45). In this study, we extend this observation. We found that an activation domain capable of inducing at least partial cell cycle arrest can be formed by activation domain 1 plus activation domain 2, activation domain 1 plus the proline-rich domain, or activation domain 2 plus the proline-rich domain (Table I). When two of the three domains, i.e. activation domain 1, activation domain 2, and the proline-rich domain, become dysfunctional, the activity in cell cycle arrest is abrogated (Table I). It should be mentioned that p53-(AD1 Ϫ ) is defective in inducing cell cycle arrest although two functional domains, i.e. activation domain 2 and the proline-rich domain are still intact (22). However, when part or all of the residues for activation domain 1 are deleted, as in p53-(⌬1-23) and p53-(⌬AD1), the ability to induce cell cycle arrest is retained. This suggests that the presence of the mutated activation domain 1 may mask the activity of, or inhibit the interaction of, a potential co-activator (or an adaptor) with the activation domain formed by activation do-main 2 and the proline-rich domain necessary for transactivation or transrepression.
The search for mediators of p53-dependent cell cycle arrest has identified many cellular p53 target genes (1,4,46). p21 cip1/waf1 , a well characterized cyclin-dependent kinase inhibitor, can mediate cell cycle arrest in G 1 when overexpressed (22,(47)(48)(49)(50)(51). Previous studies have shown that p53-(AD1 Ϫ ), which is deficient in inducing p21, is incapable of inducing arrest in G 1 , consistent with the hypothesis that p21 plays an important role in mediating p53-dependent arrest in G 1 (22,40). In this study, we found that p53-(AD2 Ϫ ) is extremely active in inducing arrest in G 1 , suggesting that activation domain 1, but not activation domain 2, plays an important role in inducing cell cycle arrest. However, p21 is only slightly induced by p53-(AD2 Ϫ ) (Fig. 1A). Because p53-(AD1 Ϫ AD2 Ϫ ), which is deficient in both activation domain 1 and activation domain 2, is inert in inducing cell cycle arrest (9 -11), this suggests that a gene(s) responsible for arrest by p53-(AD2 Ϫ ) must be induced. This is not surprising because DNA damage-induced G 1 arrest is delayed but not abolished in p21-null fibroblasts from p21deficient mice (52,53). Therefore, the cell line that inducibly expresses p53-(AD2 Ϫ ) can be used to identify other novel gene(s) responsible for G 1 arrest.
Previously, several studies have shown that the p53 protein can be cleaved by cellular proteases in cells treated with DNA damaging agents, which leads to formation of several smaller polypeptides with molecular masses ranging from 35-50 kDa (54 -58). In addition, the cleavage of p53 is concomitant with the  [11][12][13][14][15][16]. Antibodies used were as described in the legend to Fig. 1.   FIG. 6. Regulation of p21 and BAX by p53 mutants. A Northern blot was prepared using total RNAs isolated from non-induced cells (Ϫ) or cells induced for 24 h to express wild-type p53 or various p53 mutants as shown above the blot (ϩ). The blot was probed with cDNAs derived from the p21, BAX, and glyceraldehyde-3-phosphate dehydrogenase genes, respectively. After normalization to the amount of glyceraldehyde-3-phosphate dehydrogenase transcripts, the levels of induction by wild-type p53 or various p53 mutants were quantified by phosphorimager and are shown below the blot. p53 Functional Domains for Apoptosis 39933 onset of apoptosis in cells treated with DNA damaging agents, suggesting that the cleaved p53 polypeptides are potent in p53 activity and may participate in the apoptotic process (58). Interestingly, one of the cleaved p53 polypeptides, p50, is p53-(⌬N23), which lacks the N-terminal 23 residues (58). We have shown previously that p53-(⌬N23) is active in inducing cell cycle arrest and apoptosis (10). Thus, the cellular machinery can generate an active but smaller p53 polypeptide that would not be subject to negative regulation by MDM2 (59 -63). It is not clear whether p53-(⌬AD1⌬BD) is an in vivo cleavage product of p53. However, because p53-(⌬AD1⌬BD) lacks the MDM2 binding site, it would not be subjected to the negative regulation by MDM2. Thus, p53-(⌬AD1⌬BD) represents a small but potent, apoptosis-inducing form of p53. Recent clinical trials have shown that adenoviruses expressing p53 are effective in treating some advanced forms of human cancers (64,65). We suggest that p53-(⌬AD1⌬BD) is a good candidate to replace the larger, unwieldy wild-type p53 in cancer gene therapy.


extfieldsofstudy: Medicine Biology

text:
Ionizing Radiation and Bacterial Challenge Alter Splenic Cytokine Gene Expression

granulocyte-macrophage colony-stimulating factor; IL-1ß, interleukin-1ß; IL-6, interleukin-6; TNF- , tumor necrosis factor- ; Taq, Thermus aquaticus ; RT-PCR, reverse transcription-polymerase chain reaction; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; mRNA, γ radiation/ Mouse/ Spleen/ Bacterial challenge/RT-PCR Irradiation increases susceptibility to bacterial infection. Exogenous proinflammatory cytokines can alter the response of mice to γ radiation, but the role of endogenous inflammatory cytokines after bacterial infection in irradiated animals is not known. Gene expression of hematopoietic (GM-CSF) and proinflammatory (IL-1 β , IL-6 and TNF- α ) cytokines were examined in spleens of B6D2F1/J female mice after irradiation alone (1.0- and 7.0-Gy), and after irradiation followed by Klebsiella pneumoniae s.c. challenge 4 days postirradiation by using the reverse transcription-polymerase chain reaction (RT-PCR) and Southern blot hybridization. At 4, 8, and 24 h after bacterial challenge in 7.0-Gy-irradiated mice, GM-CSF mRNA increased (p < 0.05). TNF- α mRNA in irradiated mice were slightly decreased, whereas after bacterial challenge, TNF- α mRNA elevated at 30 h in 7.0-Gy-irradiated mice; at 4, and 8 h in 1.0-Gy-irradiated mice, and at 1 h in sham-irradiated mice (p < 0.05). IL-6 mRNA displayed a biphasic response in 7.0-Gy-irradiated mice, and, after bacterial challenge, in both irradiated mice (1.0– and 7.0-Gy) and sham-irradiated mice. IL-1 β mRNA remained at or below normal for 8 h and increased at 24 h after bacterial challenge on day 4 in 7.0-Gy-irradiated mice. These results indicate that sublethal gamma radiation alters the patterns of the hematopoietic and proinflammatory cytokine responses to bacterial challenge in vivo . Consequently, treatment protocols may need to take into account changes in cytokine gene responses to resolve infection after irradiation.

INTRODUCTION
It is well established that infection is a major cause of death after whole-body ionizing radiation injury 1) and that endogenous and exogenous cytokines play a major role in controlling the host's responses to radiation and infection 2) . Some cytokines are produced constitutively, but most are produced by induction 3) . Cytokines are produced by several different cell types, including monocytes/macrophages, endothelial cells, T-lymphocytes, fibroblasts, and polymorphonuclear leukocytes 4) . The first line of defense against bacterial infection involves the initiation of an inflammatory response and the release of mediators such as oxygen radicals and proinflammatory cytokines from activated macrophages and granulocytes 5) . The role of endogenous cytokines in response to radiation and bacterial infection is not clear, but administration of neutralizing antibodies against cytokines in irradiated mice has indicated that interleukin (IL)-1 and tumor necrosis factor (TNF) in blood and tissue contribute to radioresistance in mice 6) . Further, several investigators demonstrated that endogenous production of IL-6 and TNF-α is a critical component of antibacterial responses 7,8) . Also, major trauma and sepsis may lead to overproduction of TNF, IL-1, and IL-6, which may have deleterious actions on metabolic functions and hemodynamic stability 9) . Neutralizing antibodies to IL-6 and TNF-α attenuate deleterious consequences of Gram-negative sepsis caused by bacterial endotoxin 10) . Thus, an understanding of the patterns of cytokine production in a number of experimental and clinical situations could contribute to development of improved treatment protocols. Monitoring of cytokines could correlate with efficacy of therapy.
The purpose of this study was to investigate the change in expression of a single hematopoietic cytokine gene, granulocyte-macrophage colony-stimulating factor (GM-CSF), and three proinflammatory cytokine genes (IL-1β , TNF-α, and IL-6) in mouse spleen after irradiation and during bacterial challenge after sublethal gamma radiation. The spleen is a hematopoietic organ in mice. The hypothesis tested was that the gene expression of hematopoietic and proinflammatory cytokines in response to an infectious challenge is both qualitatively and quantitatively different in irradiated and sham-irradiated mice. Little is known about cytokine gene expression in vital hematopoietic organs after irradiation and subsequent bacterial challenge. In this study we used the reverse transcription-polymerase chain reaction (RT-PCR) and Southern blot hybridization techniques to determine cytokine mRNA both in mice after irradiation alone and in mice that were irradiated and then challenged with K. pneumoniae.

Mice and irradiation conditions
All animal experiments were approved by the institutional animal care and use committee. Research was conducted according to the principles enunciated in the Guide for the Care and Use of Laboratory Animals prepared by the Institute of Laboratory Animal Resources, National Research Council 11) . B6D2F1/J female mice were purchased from Jackson Laboratories (Bar Harbor, ME) and weighed approximately 20 g when irradiated in our 60 Co gamma radiation facility. Irradiation was performed bilaterally delivering total absorbed doses of 1.0 Gy and 7.0 Gy at a dose-rate of 0.4 Gy/min. In this strain of mouse, 1.0 Gy is a low sublethal dose, while 7.0 Gy is a high sublethal dose 12) . Sham-irradiated mice were treated similarly to irradiated mice but were not irradiated. Dosimetry was performed as described previously 12,13) .

Bacterial challenge
K. pneumoniae serotype 5 AFRRI 7 was obtained from a clinical isolate and prepared as previously described 14) . Four days after irradiation, mice were lightly anesthetized with methoxyflurane, and challenged s.c. with a number of K. pneumoniae that was an approximate LD 85/30 for mice at each dose of gamma radiation, i.e., 7.0-Gy-irradiated mice were given 1.8 × 10 2 colony forming units (CFU)/0.2 ml; 1.0-Gy-irradiated mice, 4.2 × 10 6 CFU/0.2 ml ; and sham-irradiated mice, 7.5 × 10 6 CFU/0.2 ml. Four days after irradiation, at the time of challenge with bacteria, peripheral blood and bone marrow cellular elements are maximally depressed 12) . The challenge doses of bacteria used assured an equivalent survival response of mice regardless of the dose of radiation.

LD 50/30 K. pneumoniae in irradiated mice
Five sets of 16 mice each were either sham-irradiated (0 Gy) or given 1.0, or 7.0 Gy radiation. Four days after irradiation, each set of 16 mice was challenged s.c. with an appropriate ten-fold serial dilution of K. pneumoniae. Survival was recorded for 30 days after bacterial challenge. A probit analysis of mortality provided the lethal dose of K. pneumoniae for 50% of mice (LD 50/30 ).

Experimental design and Statistics
Initial studies were carried out in which each specific cytokine mRNA was evaluated in the spleens of mice each 1, 4, 7, 10, 14, and 17 days after 1.0-Gy-and 7.0-Gy-irradiation, respectively. In addition in this series of studies, two sham-irradiated mice, referred to as sham I controls, were evaluated at each time point. The data from sham-irradiated mice (n = 6 -12) were pooled and are represented in the figures as a straight line. Subsequent studies were performed in which each specific cytokine mRNA was evaluated in 5 spleens each from 1.0-Gyand 7.0-Gy-irradiated mice and sham-irradiated mice at 1,4,8,24,30, and 48 h after bacterial challenge, respectively. In this series of studies, sham-irradiated, non infected mice are referred to as sham II controls. As in the initial studies, two sham-irradiated mice were evaluated at all sampling times and data from the sham-irradiated mice were pooled. Comparisons with sham I controls at each time postirradiation and with sham II controls were made using Student's t-test. In the second series of experiments comparisons were also made between irradiated-infected mice and sham-irradiated-infected mice. Transformations of the data, where needed, were made to achieve homogeneous variance. The y-axis scales in the figures reflect these transformations. Statistical analyses were done using Systat 7 (SPSS Inc., Chicago, IL). The specificity of cytokine mRNA was determined by the ratio of mRNA for each cytokine to mRNA for the housekeeping gene GAPDH for each animal. The ratio of cytokine mRNA to GAPDH mRNA in normal controls did not vary significantly (p > 0.05) between experiments. Therefore, this ratio was used as the reference point within each experiment.

RNA preparation and RT-PCR
Whole spleens were homogenized in 2 ml of RNA STAT-60 (Tel-Test Inc., Friendswood, TX) with a polytron homogenizer (Brinkmann Instruments, Inc., Westbury, NY). Homogenized spleens were immediately frozen in liquid nitrogen and kept in a -70°C freezer until mRNA analysis. Total RNA was extracted according to the manufacturer's protocol as modified by Chomczynski and Sacchi 15) . RNA concentrations were determined spectrophotometrically at 260 nm (Beckman DU-65). Total RNA for each sample was characterized by ethidium bromide-staining of agarose gels to ensure use of intact total RNA only. Intact total RNA was reverse transcribed into complementary deoxyribonucleic acid (cDNA). The final volume of 2.5 µl reverse transcription reaction mixture contained 0.5 µg total RNA, 0. Island, NY). The reaction was carried out at 37°C for 1 h, stopped at 90°C for 10 min and then chilled on ice for 10 min. The synthesized cDNA was used as a template for the PCR. Primers for GM-CSF, IL-1β , IL-6 and TNF-α were purchased from Clontech Laboratories, Inc., Palo Alto, CA, and GAPDH was purchased from Synthetic Genetics, San Diego, CA. The sequence of primers and expected sizes of each cytokine are as follows: GM-CSF(368 bp) 5'-primer 5'TGTGGTCTACAGCCTCTCAGCAC3' 3'-primer 5'CAAAGGGGATATCAGTCAGAAAGGT3' IL-1β (563 bp) 5'-primer 5'ATGGCAACTGTTCCTGAACTCAACT3' 3'-primer 5'CAGGACAGGTATAGATTCTTTCCTTT3' IL-6 (638 bp) 5'-primer 5'ATGAAGTTCCTCTCTGCAAGAGACT3' 3'-primer 5'CACTAGGTTTGCCGAGTAGATCTC3' TNF-α (354 bp) 5'-primer 5'TTCTGTCTACTGAACTTCGGGGTGATCGGTCC3' 3'-primer 5'GTATGAGATAGCAAATCGGCTGACGGTGTGGG3' The reaction mixture for PCR contained 5 µl cDNA template from the RT reaction, 10 mM Tris-HCl (pH 8.3), 50mM KCl, 1.5 mM MgCl 2 , 0.8 mM each of dATP, dCTP, dGTP, and dTTP, 1.0 µM 5 ' -primer, 1.0 µM 3 ' -primer, and 1.25 U Taq ( Thermus aquaticus) DNA polymerase. PCR was performed with a DNA Thermal Cycler (Perkin Elmer-Cetus, Norwalk, CT). PCR reactions were carried out on cDNA templates prepared in the absence of reverse transcriptase as negative controls. A positive control for each cytokine was also included in all PCR reactions. Cycle number was optimized individually for each cytokine as described by Gause and Adamovicz 16) . The amplified products were characterized in 1% agarose gel electrophoresis, transferred onto a Nytran nylon membrane (Schleicher and Schull, Keene, NY), and hybridized with the respective fluorescein-labeled GM-CSF, IL-6, TNF-α, GAPDH plasmid cDNA probe, and IL-1β oligonucleotide probe. The probes for IL-6, TNF-α, and GAPDH were prepared as described previously 17) . The GM-CSF probe consisted of a 0.8-kb insert cloned into the EcoRI site of pXM (Genetics Institute, Cambridge, MA). Probes for GM-CSF, IL-6, TNF-α and GAPDH were labeled with a random priming commercial kit (Amersham, Arlington Heights, IL). The probe for IL-1β was labeled using an ECL 3'-oligolabelling commercial kit (Amersham, Arlington Heights, IL). Hybridization, stringency washes, and signal detection were carried out according to the manufacturer's instructions. Fluorographs were prepared at room temperature on Kodak XAR film. The densitometric volume of the PCR product band of each sample was measured using a scanning laser densitometer (Molecular  Dynamics, Sunnyvale, CA). Semiquantitation was based on relative cytokine mRNA differences between treatment and control samples which were normalized to its GAPDH value.

Relative GM-CSF, IL-6, and TNF-α mRNA in spleens after irradiation
Relative GM-CSF mRNA responses increased (p > 0.05) up to 10 days after 7.0 Gy (Fig. 1A) when compared to sham I controls. No significant changes in GM-CSF mRNA was observed 10, 14 and 17 days after 1.0-Gy-and 7.0-Gy-irradiation. Data are expressed as the ratio of IL-6 to GAPDH and represent the mean ± SEM from three to five mice per treatment group and twelve mice for sham I controls group. *p < 0.05 and **p < 0.01 with respect to the sham I control value. The time period between vertical dashed lines is equivalent to that in Fig. 6A. Relative IL-6 mRNA responses were significantly increased (p < 0.05) above sham I control values (n = 12) 1, 4, and 10 days only after 7.0 Gy. The increases in IL-6 mRNA occurred in a biphasic pattern, with peaks at days 1 and 10 ( Fig. 2A). A representative fluorograph of RT-PCR-amplified mRNA transcripts of IL-6 and GAPDH in spleens of sham I controls and 7.0-Gy-irradiated mice at days 1, 4 and 10 is shown in Fig. 2 B.    Relative TNF-α mRNA of irradiated mice were not different from sham I control values on the days tested (Fig. 3A). A representative fluorograph of RT-PCR-amplified mRNA transcripts of TNF-α and GAPDH in spleens of sham I controls and 7.0-Gy-irradiated mice at days 1, 4, and 10 is shown in Fig. 3B. The change in relative IL-1β mRNA was not determined in this experimental set. In Figs. 1A, 2A, 3A the time period of days 4 to 6 after radiation is inicated by dashed vertical lines and is equivalent to the 0 -48 h time period after    similar to those of sham-irradiated-infected mice, except that GM-CSF mRNA in sham-irradiated-infected mice increased (p < 0.05) at 1 hr over sham II controls (n = 6) (Fig. 4A). A representative fluorograph of RT-PCR-amplified mRNA transcripts of GM-CSF and GAPDH in  Data are expressed as the ratio of TNF-α to GAPDH and represent the mean ± SEM from three to five mice per treatment group and six mice for sham II controls group. *p < 0.05 and **p < 0.01 with respect to sham II control value. Fig. 4B. Relative IL-1β mRNA in 1.0-Gy-and sham-irradiated mice decreased below sham II controls at 1 h after bacterial challenge, then increased to a higher level at 4 h and then decreased again. Relative IL-1β mRNA in 7.0 Gy-irradiated-infected mice were different from those in 1.0-Gy-and sham-irradiated-infected mice. Relative IL-1β mRNA in the 1.0-Gy-and sham-irradiated-infected mice did not differ significantly across time, on the other hand in 7.0-Gy-irradiated-infected mice, relative IL-1β mRNA rose at 24 h and remained higher than those in sham II controls through 48 h. Significant differences (p < 0.05) from sham II controls (n = 6) were observed for 1.0-Gy-irradiated-infected mice at 24 h and sham-irradiatedinfected mice at 4 h (Fig. 5A). A representative fluorograph of RT-PCR-amplified mRNA transcripts of IL-1β and GAPDH in spleens of sham II controls, sham-irradiated-infected mice at 4 h and 7.0-Gy-irradiated-infected mice at 24, 30 and 48 h is shown in Fig. 5B.

spleens of sham II controls and 7.0-Gy-irradiated-infected mice at 1, 4, 8 and 24 h is shown in
The increases in relative IL-6 mRNA after K. pneumoniae challenge occurred in a biphasic pattern (Fig. 6A) Fig. 6B.
Relative TNF-α mRNA in 7.0-Gy-irradiated-infected mice were initially different from those of 1.0-Gy-and sham-irradiated-infected mice (Fig. 7A). Relative TNF-α mRNA in the 7.0-Gy-irradiated-infected mice decreased slightly below the sham II control level at 1 and 4 h, while responses in 1.0-Gy-and sham-irradiated-infected mice were initially elevated at 1, 4 and 8 h, then declined to normal levels. There were no significant differences in responses between bacteria-challenged mice that were given 1.0 Gy and sham-irradiation. Significant differences (p < 0.05) from the sham II controls (n = 6) were observed in K. pneumoniae-challenged mice after 7.0 Gy at 30 h (1.6-fold increase); after 1.0 Gy at 4 and 8 h (2.0-and 2.1fold increases); and after sham-irradiation at 1 h (2.1-fold increase). A representative fluorograph of RT-PCR-amplified mRNA transcripts of TNF-α and GAPDH in spleens of sham II controls, sham-irradiated-infected mice at 1 h, 1.0-Gy-irradiated-infected mice at 4 h and 8 h, and 7.0-Gy-irradiated-infected mice at 30 h is shown in Fig. 7B.

DISCUSSION
This pioneering study demonstrated changes in cytokine gene expression in spleens of mice in response to sublethal gamma radiation and bacterial infection. K. pneumoniae is a virulent pathogen that induces an impressive inflammatory response in mice 18) . The initiation, maintenance, and eventual resolution of inflammation as a response to bacterial challenge depend on the expression of a complex network of both pro-inflammatory and anti-inflammatory cytokines. Elevated levels of TNF-α, macrophage inflammatory protein-2, and IL-10 in lung have been observed during K. pneumoniae infection 19) .
In this present paper we used the RT-PCR and Southern blot hybridization techniques to detect alterations in cytokine mRNA in spleens of mice after acute sublethal irradiation and in sublethally irradiated mice given a bacterial challenge. We report that the responses of se-lected cytokines to the infectious challenge varied in patterns that depended upon the dose of radiation. However, the extent of the change in mRNA differed with time among various cytokines, suggesting that the combination of radiation and bacterial challenge produced different effects on cytokine mRNA transcription in the mouse.
Our studies showed a consistent trend for slightly elevated relative GM-CSF mRNA at days 4 and 7 after 7.0 Gy (Fig. 1A). During the equivalent period at 4, 8 and 24 hr in 7.0 Gyirradiated-infected mice (time points located between vertical dashed lines in Fig. 1A), GM-CSF mRNA responses were significantly higher than those in the sham II control values (p < 0.05) (Fig. 4A). However, GM-CSF mRNA in sham-irradiated mice or mice given 1.0 Gy were similar at 4 h or longer following bacterial challenge. Bacterial challenge therefore appears to further increase the GM-CSF response induced by radiation alone. Along these lines, it was shown that GM-CSF-deficient mice had a propensity to develop lung and soft-tissue infection, and also had a significantly impaired long-term survival 20) . However, Basu et al 21) showed that GM-CSF-deficient mice (a mix of C57BL/6 and 129/OLA strains) had a survival advantage with 58% of animals surviving at 50 hr as compared with 0% in the wild type following 500 µg LPS i.v. treatment on day zero. Along these lines, we have shown repeatedly that 7.0-Gy-irradiated mice do not survive challenge with K. pneumoniae 22) .
Several investigators detected an elevated accumulation of IL-1β in whole spleen after exposure to ionizing radiation. Nemoto et al 23) demonstrated that IL-1β mRNA increased at 5-7 days after 3 Gy X rays in C3H/He male mouse. The increase of IL-1β at 3 Gy is less than 2-3 times the level at time zero. In our laboratory, the LD 50/30 for B6D2F1/J female mouse is 9.0 Gy, whereas the LD 50/30 for C3H/He male mice is 7.32 Gy. Further, X rays share a higher relative biological effect than 60 Co gamma rays. Ishihara et al 24) showed that immediate accumulation of IL-1β mRNA was found 30 min after the initiation of supra-lethal irradiation with 18.5 Gy X rays in C3H/HeN mouse and the increased level of message was not decreased until 4 h. In our results, relative IL-1β mRNA in 7.0-Gy-irradiated-infected mice decreased during the first day, then increased during the second day, but were elevated in sham-irradiated-infected mice during the first 4 h (Fig. 5A). This finding suggests that sublethal irradiation inhibits the increase of IL-1β gene transcription and leads directly or indirectly to increased susceptibility to infection. Therefore, IL-1β may have an important role in promoting innate resistance to bacterial infection in irradiated mice. In support of this, Peterson et al 22) found that two bacterial derived immunomodulators, synthetic trehalose dicorynomycolate (S-TDCM) and an extract from Serratia marcescens (Sm-BRM) increased resistance to K. pneumoniae infection and also elevated gene expression of IL-1β in the spleen for 5 days after 7.0 Gy γ radiation. Along these lines, prophylactic administration of a low dose of recombinant IL-1β to mice 24 h before Pseudomonas aeruginosa infection increased the natural resistance to infection in granulocytopenic mice 25) .
Changes in relative IL-6 mRNA occurred in a biphasic manner in different treatment groups at different times (Fig. 6A). That is, the biphasic pattern became more delayed with increasing dose of radiation. Relative IL-6 mRNA did not significantly decrease during the first hour after infection, but increased in sham-irradiated-infected mice at 4 h. Relative IL-6 mRNA decreased 8 h after bacterial challenge, then increased the next day in 1.0-Gy-and 7.0-Gy-irradiated-infected mice, but not in sham-irradiated-infected mice. Rapid alteration of IL-6 mRNA after infection is possible. Bacteria can induce a particular cytokine network, or alter an existing one, by stimulating the release of a variety of cytokines from host cells 26) . Waage et al 27) demonstrated that after administration of meningococcal (Neisseria meningitidis) LPS to the subarachnoid space of rabbits, IL-6 concentration rose in the cerebrospinal fluid within 2 h, peaked after 3 h and declined to baseline after 9 h. Rola-Pleszczynski et al 28) demonstrated that Leukotriene (LTB) stimulated IL-6 production and that underlying mechanisms involved both increased IL-6 gene transcription and message stabilization. This may constitute an important mechanism through which rapidly produced mediators may modulate the subsequent production of regulatory or growth-promoting cytokines. Along these lines, Peterson et al 22) observed that S-TDCM and Sm-BRM treatments of 7.0-Gy-irradiated mice elevated splenic IL-6 mRNA after 3 days. Elevation of IL-6 protein was observed in spleen cells from Mycobacterium avium-infected C57BL/6 mice at 2 weeks 29) and in the circulation and spleens of mice infected with Staphylococcus aureus at 24 h 30) . Some investigators have considered persistently elevated IL-6 to predict mortality 31) . However, Patchen et al 32) showed that therapeutically administered IL-6 can effectively accelerate multilineage hematopoietic recovery following radiation-induced hematopoietic injury. Also, Neta et al 33) demonstrated that interaction of IL-6 with IL-1 and TNF is a prerequisite for protection from radiation lethality.
In our studies relative TNF-α mRNA remained essentially normal in irradiated mice (Fig. 3A). In contrast, relative TNF-α mRNA was slightly below control values within 4 hr after bacterial challenge in 7.0-Gy-irradiated-mice, whereas they were immediately elevated in 1.0-Gy-and sham-irradiated-infected mice (Fig. 7A). K. pneumoniae challenge promoted elevation of relative TNF-α mRNA in 1.0-Gy-irradiated-infected mice at 1-8 h and at 1-4 h in sham-irradiated-infected mice. Further, Peterson et al 22) found that TNF-α mRNA remained below normal levels 3-7 days after 7.0 Gy. Thus, elevation of TNF-α mRNA in 7.0-Gy-irradiated mice by K. pneumoniae appears to be delayed a few hours, after an initial depression. Increases in TNF-α mRNA after bacterial infection has been associated with septic shock and pathogenesis of infectious diseases, but, if regulated, TNF-α may provide the host a beneficial response to infection 34) .
Peterson et al 22) also demonstrated that S-TDCM and Sm-BRM enhanced survival from K. pneumoniae challenge in irradiated mice. In mice treated with these immunomodulators, elevated IL-1β and IL-6 mRNA together with a low TNF-α mRNA at the time of bacterial challenge appeared to be associated with survival from subsequent bacterial infection in irradiated mice. In contrast, our results showed that IL-1β, IL-6 and TNF-α mRNA remained near control values 1-8 h after K. pneumoniae challenge in 7.0-Gy-irradiated mice. It would be appropriate, therefore, to examine the cytokine response that promotes resistance to infection in irradiated mice by determining cytokine gene expression in spleens of mice that are irradiated, treated with a nonspecific immunomodulator, known to enhance survival from infection, and then challenged with bacteria.
The changes we measured in proinflammatory cytokine gene expression in irradiated mice indicate a possible relationship with increasing susceptibility to infection. A slightly delayed and prolonged increase in transcription of TNF-α and GM-CSF genes associated with biphasic responses of IL-6 mRNA indicates a poor prognosis for survival in acutely irradiated and infected animals. In contrast, elevated IL-1 mRNA coupled with a suppressed TNF-α mRNA response indicates a favorable prognosis due to enhanced resistance to infection. This concept needs further testing.
The effect of radiation on cells may be due to either direct interaction of the radiation with target molecules or indirectly through the generation of free radicals and oxidizing and reducing chemical species. Molecular damage caused by radiation is expressed through alteration of biochemical processes and is amplified by biological mediators 36) . Cytokines, one type of mediators, are signaling molecules which can behave as classic endocrine hormones, but are better known for their autocrine and paracrine behavior, acting as integrating signals over short cellular distances. It is now realized that cytokines rarely, if ever, act in isolation but rather induce or inhibit other cytokines, thus, creating a population, or network, of cytokines to which cells respond 26) . Ibuki et al 37) reported that the activation of macrophages by radiation is caused by interaction with neighboring cells, such as lymphocytes, and by paracrine induction of certain cytokines which is initiated by the small amount of IL-1β released by irradiated macrophages. In the current paradigm, cytokine induction as a protective or pathological mechanism, is a direct response to the presence of infectious microorganisms.
It is now clear that bacteria contain and produce a large number of diverse molecules, which can selectively induce the synthesis of both pro-inflammatory and immunomodulatory/anti-inflammatory cytokines 38) . Cytokines play an essential role in orchestrating the host's response to radiation, infection, trauma, and other forms of injury. No work has been reported on gene expression and their associated proteins in irradiated animals with an induced infection. In previous studies, administration of recombinant cytokines as well as neutralizing antibodies to cytokines, established that IL-1 synergizes with TNF, CSFs, and IL-6 to protect mice from hematopoietic failure. In contrast, cytokines such as transforming growth factor (TGF)-β and interferon-gamma (IFN-γ ) sensitize untreated mice to radiation lethality 6) . Thus, individual cytokines can be either beneficial or detrimental to the host's recovery from damage due to ionizing radiation. An understanding of the signal transmitted to target cells by one or more cytokines and the context in which they operate will provide insight into the regulation of hematopoiesis, inflammation, wound closure (i.e., healing), and sepsis in irradiated animals. A better understanding may also lead to improved strategies for management of infection 39) . Studies of bacterial infection, particularly sepsis, require intricate physiological interactions among cells, tissues and organs. Resistance to infection depends upon a complex network of interacting signaling systems that invlove the innate and adaptive immune systems, nervous system, and endocrine system. The physiology of these interacting networks is altered by radiation. The effects and responses, therefore, must be examined in whole, integrated animals, rather than limited cell or tissue cultures. Cell culture can be used as an analytic tool, but cell cultures cannot substitute for the complex interactions in an intact animal that provides the integrated biochemical, physiological, cellular, microbial, and multi-organ interactions necessary to perform these studies. The extrapolation of data from in vitro cell or tissue cultures to accurately describe even a single facet of the immune response of the whole animal is difficult in studies designed to determine the effects of various treatments, such as ionizing radiation or drugs, on infection. Only in living, whole animals can the interaction of all the functioning components of the immune system be examined. It is therefore necessary to perform studies of the combined effects of ionizing radiation and bacteria in living expeimental animals.
The data presented here provide impetus for subsequent studies on the role of proinflammatory cytokines in the pathogenesis of bacterial challenge in irradiated mice. We are concurrently determining cytokine proteins in spleen and plasma by using the ELISA method in another aspect of this study. These studies could lead to improved evaluation of the status of animals during the course of infection and efficacy of therapeutic interventions. Determining optimal cytokine responses for resistance to bacterial infection will aid in the development of potential therapeutic strategies for infection after radiation injury. mens, M. E. Greenville for editorial assistance and M.A. Behme for graphic assistance. This work was supported by the Armed Forces Radiobiology Research Institute under work unit 09505.


extfieldsofstudy: Biology Medicine

text:
Conformational Requirements for Glycoprotein Reglucosylation in the Endoplasmic Reticulum

Newly synthesized glycoproteins interact during folding and quality control in the ER with calnexin and calreticulin, two lectins specific for monoglucosylated oligosaccharides. Binding and release are regulated by two enzymes, glucosidase II and UDP-Glc:glycoprotein:glycosyltransferase (GT), which cyclically remove and reattach the essential glucose residues on the N-linked oligosaccharides. GT acts as a folding sensor in the cycle, selectively reglucosylating incompletely folded glycoproteins and promoting binding of its substrates to the lectins. To investigate how nonnative protein conformations are recognized and directed to this unique chaperone system, we analyzed the interaction of GT with a series of model substrates with well defined conformations derived from RNaseB. We found that conformations with slight perturbations were not reglucosylated by GT. In contrast, a partially structured nonnative form was efficiently recognized by the enzyme. When this form was converted back to a nativelike state, concomitant loss of recognition by GT occurred, reproducing the reglucosylation conditions observed in vivo with isolated components. Moreover, fully unfolded conformers were poorly recognized. The results indicated that GT is able to distinguish between different nonnative conformations with a distinct preference for partially structured conformers. The findings suggest that discrete populations of nonnative conformations are selectively reglucosylated to participate in the calnexin/calreticulin chaperone pathway.

Introduction
The ER provides the folding and assembly environment for a multitude of soluble and membrane-bound proteins destined to diverse intra-and extracellular locations. Protein maturation in this organelle involves translocation, co-and posttranslational modifications, disulfide bond formation, and chaperone-assisted folding. In addition, newly synthesized proteins are subjected to quality control, which restricts forward transport to correctly folded and oligomerized conformers (Ellgard et al., 1999). Failing to reach the native conformations, proteins are targeted for selective degradation (Klausner and Sitia, 1990).
Of the systems that assist and monitor protein maturation, the calnexin/calreticulin cycle is unique in that it di-rectly depends on the structure of oligosaccharides on newly synthesized glycoproteins (Bergeron et al., 1994;Williams, 1995;Helenius et al., 1997). Calnexin and calreticulin are homologous lectins that bind to the monoglucosylated forms of N-glycans Glc 1 Man 6-9 GlcNAc 2 . This association promotes correct folding and oligomerization, supports ER retention of folding intermediates and misfolded or incompletely assembled forms, and exposes them to the glycoprotein-specific thiol oxidoreductase ERp57 (Trombetta and Helenius, 1998).
The monoglucosylated oligosaccharides central to this chaperone system are generated in two ways. Initially, they arise by the removal of the two outermost glucoses by the sequential action of glucosidases I and II, a rapid process which is independent of the folding status of the glycoproteins (Kornfeld and Kornfeld, 1985). Monoglucosylated glycans are also generated by an ER lumenal enzyme, the UDP-glc:glycoprotein:glucosyltrans-ferase (GT), 1 which transfers a glucose residue from UDP-Glc to fully deglucosylated glycans, reestablishing the glucose-␣ ( 1,3)-mannose glycosidic bond recognized by calnexin and calreticulin. The opposing actions of glucosidase II and GT establish a deglucosylation/reglucosylation cycle that drives a cycle of calnexin and calreticulin binding and release (Hammond et al., 1993).
GT is a soluble glycoprotein of 170 kD with an ER localization signal ubiquitously expressed in the ER of most cell types, tissues, and species (Trombetta and Parodi, 1992;Parker et al., 1995;Fernandez et al., 1996). Glycoprotein reglucosylation is a selective reaction that only occurs for incompletely folded proteins (Suh et al., 1989;Sousa et al., 1992;Trombetta and Parodi, 1992;Fernandez et al., 1994;Hebert et al., 1995;Parker et al., 1995;Van Leeuwen and Kearse, 1997;Wada et al., 1997;Cannon and Helenuis, 1999;Labriola et al., 1999). While the in vitro studies do not rule out the possibility that it cooperates with other ER factors in vivo, they strongly suggest that the conformational sensor function is primarily, if not exclusively, performed by GT itself. Work by Parodi and coworkers has shown that to be reglucosylated, i.e., glycans have to be linked to the polypeptide backbone of a protein with a nonnative conformation, and the innermost GlcNAc bound to the Asn is a key element in the recognition process (Sousa et al., 1992;Trombetta and Parodi, 1992;Sousa and Parodi, 1995). It has been proposed that GT senses the exposure of hydrophobic regions normally buried in native conformations, in a way similar to HSP70 and other molecular chaperones (Sousa and Parodi, 1995).
However, the mechanism whereby GT discriminates between native and nonnative protein conformations remains largely unexplored. This is in part due to the experimental challenge imposed by the nonnative nature of the GT substrates. Once proteins depart from their native conformation, they tend to aggregate and behave in ways that are difficult to control and characterize. To analyze the conformational requirements for glycoprotein reglucosylation we generated a panel of defined conformers of pancreatic RNaseB and studied their interaction with GT. Using monomeric variants of RNaseB as model substrates, we found that exposure of an otherwise buried GlcNAc-Asn bond, in a slightly perturbed conformer or in a completely unfolded polypeptide chain, was not a sufficient condition for reglucosylation. On the other hand, a partially structured conformation was better recognized by GT than a completely unfolded form, indicating that not all nonnative conformations can be equally reglucosylated by GT, which showed a preference for structured folding intermediates. These results show that distinct nonnative protein conformations are selectively reglucosylated to participate in the calnexin/calreticulin pathway.

Proteins
RNaseB (Sigma R5870) and RNaseA (Sigma R5125) were purified by chromatography on a 2.5 ϫ 60-cm column of BioRex70 (BioRad) by isocratic elution at 1 ml/min with 0.2 M sodium phosphate pH 6.5. Prepara-tive digestions with EndoH (New England Biolabs 702S) or ␣ -mannosidase (Sigma M7257) were performed in 0.1 M sodium acetate buffer, pH 5.0 at 20 Њ C for 24 h. Digestions with subtilisin (Sigma P-4789) were performed at a protease/substrate ratio of 1:750 and a protein concentration of 10-50 mg/ml in 0.1 M Tris-HCl, pH 8.0. After 48 h on ice, pH was brought to 2.5 with ice-cold 1 N HCl. The acidified samples were incubated for 1 h on ice and the pH was brought back to 6.5 with ice-cold 5 N NH 3 . Samples were purified on BioRex70 as described above.
For preparation of RNaseBS-Prot, 1 ml of glacial acetic acid was added to the 1 ml of RNaseBS on ice. The acidified samples were run on Sephadex G-50 medium (Pharmacia; 1.5 ϫ 45 cm), freshly equilibrated and run in 50% acetic acid at 4 Њ C. RNaseBS-Prot eluted in the void was concentrated in Speed Vac to 0.5 ml and the buffer was exchanged by gel filtration into 50 mM NaCl and 20 mM Hepes, pH 7.5. To regenerate RNaseBS Ј and RNaseBS ЈЈ , freshly prepared RNaseBS-Prot was incubated with a 5-M excess of S-peptides in 50 mM NaCl and 20 mM Hepes, pH 7.5, and purified in BioRex70 as described.
To prepare reduced and alkylated RNaseB, 10 mg of protein was reduced in 1 ml of 8 M urea, 10 mM DTT, and 0.1 M Tris-HCl, pH 8.0. After 30 min at 37 Њ C, the alkylating agents were added at a final concentration of 80 mM, and samples were incubated for a further 30 min at 37 Њ C, after which the samples were desalted into 50 mM NaCl and 20 mM Hepes, pH 7.5.
In all cases, the purity was analyzed by SDS-PAGE and by analytical chromatography on MonoS 5/5 (Pharmacia) in 20 mM Hepes, pH 7.5, eluted with a gradient from 0 to 150 mM NaCl during 45 min at 1 ml/min. Different ribonuclease samples were stored at Ϫ 80 Њ C in 0.2 M sodium phosphate, pH 6.5. To perform the assays, they were exchanged into 50 mM NaCl, 20 mM Hepes, pH 7.5, buffer and concentrated by ultrafiltration. In some experiments, the nonnative conformers were supplemented with 20% sucrose and 0.1% octylglucoside to prevent aggregation, which did not affect the glucosylation reactions.
To analyze the sensitivity of different conformers to EndoH, N -glycanase, or trypsin, 1 g of protein was incubated with 250 U of EndoH (New England Biolabs 702), or 10 U of N -glycanase (New England Biolabs 704) or 0.01 g of trypsin (Sigma T8642) in a final volume of 10 l containing 10 mM CaCl 2 , 50 mM NaCl, 20 mM Hepes, pH 7.5. After 5-60 min at 30 or 37 Њ C, samples were analyzed by SDS-PAGE and stained with Coomasie blue R-250.

Assays
GT was purified as previously described (Trombetta and Parodi, 1992). Glucosylation reactions were performed in a final volume of 50 l containing 100-500 g of acceptor glycoproteins, 10 mM CaCl 2 , 50 mM NaCl, 20 mM Hepes, pH 7.5, 0.05-0.1 M UDP[ 3 H]Glc. Reactions were initiated by addition of 5 l of GT (0.1-0.5 g) and incubated at 30-37 Њ C for 5-10 min. For inhibition studies, 50-100 g of urea-denatured soybean agglutinin (SBA) was used as an acceptor, and ribonuclease forms were added at a one-to fivefold molar excess.
The ribonuclease activity was measured in a total volume of 140 l containing 0.5% RNA (Sigma R6625), 20 mM NaCl and 50 mM Hepes buffer, pH 7.5. After equilibration at 30 Њ C, reactions were started by the addition of 5 l of ribonuclease (140 g/ml). Aliquots of 20 l were removed at 30-s intervals from the incubations and mixed with 10 l of 25% perchloric acid and 0.8% uranylacetate. After 10 min at 20 Њ C, samples were centrifuged for 5 min at 14,000 g , 10 l of the supernatant was mixed with 300 l of water, and the absorbance was measured at 260 nm.

Generation of RNaseB Conformers
As a model substrate for GT, we selected bovine pancreatic ribonuclease B (RNaseB), a small (124 amino acids), soluble, and well characterized glycoprotein. Since it only differs from RNaseA by the presence of a single high mannose oligosaccharide on Asn34, we generated different conformers of RNaseB by adapting procedures previously described for RNaseA. We implemented a modification known as the ribonuclease S system (Richards, 1958;Richards and Vithayathil, 1959) in which the peptide bond be-tween Ala20-Ser21 is cleaved by limited proteolysis, yielding an altered but fully active ribonuclease. While similar, intact RNaseA and the cleaved RNAseS differ in certain aspects: they can be separated chromatographically (Richards and Vithayathil, 1959); RNaseS is sensitive to proteases that do not digest RNaseA (Allende and Richards, 1962), and the region near the peptide bond cleaved in RNaseS is disordered and mobile (Kim et al., 1992).
Since the peptide bond cleaved in RNaseS is located near the glycosylated Asn34, we introduced a localized perturbation in the vicinity of the glycosylation site of RNaseB by reproducing the subtilisin treatment previously described for RNaseA. Cleavage was quantitative and the resulting form, hereafter designated RNaseBS, had full ribonuclease activity (Fig. 1 C). The subtilisin cleavage was also applied to RNaseB, which had been previously digested with endoglycosidase H to remove all but the innermost GlcNac residue of the N-linked glycan, and to RNaseB treated with ␣ -mannosidase to remove the mannoses of the glycan (Fig. 1 B). RNaseA was also converted into RNaseS to serve as a nonglycosylated control.
One key feature of the RNaseS system is the possibility of reversible removal and readdition of the NH 2 -terminal fragment (the S-peptide, residues 1-20), which remains bound to the protein after cleavage by subtilisin (Richards, 1958). The protein, which is devoid of a peptide (called RNaseS-Prot) is soluble but enzymatically inactive (Richards, 1958). Since it contains all four intrachain disulfide bonds, it retains much of the native structure (Richards and Wickoff, 1971;Richards et al., 1972). We found that after removal of the NH 2 -terminal peptide from RNaseBS, the remaining core (RNaseBS-Prot) was well behaved, soluble, and monomeric (Fig. 2, C and H). Like RNaseS-Prot, it avidly bound the NH 2 -terminal peptide (residues 1-20) to generate RNaseBS Ј . It also bound a shortened version of the peptide (residues 1-15) to generate RNaseBS ЈЈ . Both reconstituted forms possessed full ribonuclease activity (Fig. 1 C).
Since the disulfide bonds are critical for the folded structure, reduction of denatured RNaseA results in an unfolded polypeptide (Sela et al., 1957). To obtain such unstructured forms, RNaseB was denatured, reduced, and alkylated with iodoacetamide or iodoacetic acid, which produced soluble products ( Fig. 1 and Fig. 2 J), or with N-ethylmaleimide or 4-vinylpyridine, resulting in insoluble forms.

Characterization of the Different Conformers of RNaseB
The procedures described above allowed us to generate a family of conformers and glycoforms, some of which are shown in Fig. 1. They were all soluble, with the exception of the NEM-and 4VP-alkylated forms. Most importantly, they were monomeric under the experimental conditions used (Fig. 2, F-J). Since reduced and alkylated RNase is monomeric (Harrington and Sela, 1959), the increased Stokes radius (Fig. 2 J) is most likely due to a less compact structure consequence of unfolding and the loss of the disulfide bonds. Thus, they could be used to analyze the role of protein conformation in their interaction with GT, without the complications derived from substrate association and aggregation.
To evaluate the local environment around the glycosylated Asn34, we used endoglycosidases as macromolecular probes. In native glycoproteins, the innermost GlcNAc residue can be secluded by the surrounding polypeptide environment, preventing access of endoglycosidases. This is the case for intact RNaseB, in which the glycan can be readily cleaved by EndoH (Tarentino et al., 1974), but is much more refractive to cleavage by N -glycanase (Taren-tino and Plummer, 1982;Tarentino et al., 1985;Rudd and Dwek, 1997), which cleaves between the Asn side chain and the first GlcNAc. We found that all the conformers generated were sensitive to EndoH (Fig. 3 A), indicating that the GlcNAc-GlcNAc bond remained accessible. However, with the exception of native RNaseB, all the other soluble conformers were readily degraded by N -glycanase (Fig. 3 B). This indicated that their Asn34 was more accessible than in native RNAseB. The environment around the glycosylation site had thus changed, even in RNaseBS, in which the S-peptide remained bound and the ribonuclease activity was unaffected.
To further characterize the conformers generated, we tested their protease sensitivity. Native RNaseB was resistant to cleavage by trypsin, but all other conformers were readily digested, indicating that they had a more relaxed structure than native RNaseB (Fig. 3 C). The same pattern of protease sensitivity was obtained with chymotrypsin (not shown). Despite the marked susceptibility to protease digestion of the different conformers, the forms expected to have ribonuclease activity were fully active on RNA (Fig. 1 C).
Ion exchange chromatography was also used to monitor the different conformers. The technique was sensitive to changes in protein conformation and could differentiate between RNaseB, RNaseBS Ј , and RNaseBS ЈЈ (Fig. 2, A-E).  It also revealed that freshly isolated RNaseBS-Prot eluted as a rather homogeneous population. However, upon storage, it slowly populated different conformers and displayed more heterogeneous elution patterns. Although heterogeneous, they could still rebind S-peptide and recover full ribonuclease activity (not shown). Therefore, for the assays described below, RNaseBS-Prot was used immediately after removal of the S-peptide and neutralization.

Interaction of the Different Conformers with GT
To study the interaction of the various conformers with GT, we quantitated their capacity to be glucosylated in vitro by the isolated glucosyltransferase. The glucosylation assays used measured the rate of incorporation of [ 3 H]Glc from UDP[ 3 H]Glc into acceptor glycoproteins under conditions of initial velocity. Fig. 4 A shows that native RNaseB and mock-treated RNaseB were not recognized as substrates by GT as expected. Moreover, the enzyme was apparently unable to detect the altered conformation of RNaseBS. On the other hand, the form without the S-peptide, RNaseBS-Prot, was glucosylated by GT. As shown in Figs. 4 A and 5, the ability of RNaseBS-Prot to act as a substrate for GT was almost completely abolished upon addition of the NH 2 -terminal S-peptide (residues 1-20) or a truncated form of it (residues 1-15) to regenerate RNaseBS Ј or RNaseBS ЈЈ , respectively. Neither peptide affected reglucosylation of other denatured glycoproteins (not shown). The loss of recognition by GT upon reconstitution of ribonuclease activity was, due to the restoration of a nativelike protein conformation upon rebinding of the S-peptides to the RNaseBS-Prot since it paralleled the stoichiometric regeneration of RNaseBS Ј or RNaseBS ЈЈ (Fig. 5).
Interestingly, the unfolded conformations presented by the soluble derivatives obtained by reduction and alkylation with iodoacetamide or iodoacetic acid were poorly recognized by GT (Fig. 4 A). The aggregation of RNaseB by reduction and alkylation with NEM or 4VP, on the other hand, converted them into much better substrates for GT (Fig. 4 A).
RNaseB forms were less efficient substrates than denatured soybean agglutinin (SBA, not shown), a glycoprotein that contains a single Man 9 GlcNAc 2 oligosaccharide. In contrast, RNaseB carries a mixture of oligosaccharide structures (Man 4-9 GlcNAc 2 ), of which only a minor fraction has the terminal mannose acceptor for the Glc residue transferred by GT (Trombetta and Parodi, 1992). Since the remaining glycoforms all contain a GlcNAc residue linked to Asn34, they have the potential to behave as competitive inhibitors in the glucosylation assay (Sousa et al., 1992). To test whether this was the case and to further evaluate the interaction of the different RNaseB forms with GT, we measured the ability of the different conformers and glycoforms to inhibit GT glucosylation of denatured SBA.
As shown in Fig. 4 B, in native RNaseB, mock-treated RNaseB, RNaseBS and RNaseS-Prot, or the EndoH-or ␣ -mannosidase-digested forms of RNaseBS did not inhibit GT. This indicated that they did not interfere with the substrate recognition site of the glucosyltransferase  . Reversible recognition of RNaseBSProt by GT. The indicated amounts of S-peptides 1-20 (solid diamonds) or 1-15 (solid circles) were added to samples containing 500 g of RNaseBS-Prot in a total volume of 45 l of 10 mM CaCl 2 , 50 mM NaCl, 20 mM Hepes, pH 7.5. After 5 min at 20ЊC, glucosylation reactions were started with 5 l containing 0.3 g of GT and 400,000 cpm of UDP[ 3 H]Glc. Reactions were incubated for 5 min at 30ЊC, stopped with 1 ml of 10% TCA, and radioactivity incorporated in TCA insoluble material quantified. Parallel incubations lacking UDP[ 3 H]Glc were used to measure ribonuclease activity (empty diamonds and circles) as described in Materials and Methods. and did not affect its interaction with denatured SBA. In contrast, all forms that had a carbohydrate bound to Asn34 on the protein backbone of RNaseBS-Prot were potent inhibitors (Fig. 4 B). Although EndoH and ␣ -mannosidase-digested RNaseBS-Prot were not glucose acceptors, they were efficient GT inhibitors, showing that sugar residues extending beyond the first Asn-bound GlcNAc had little effect on interaction with GT. The soluble reduced/alkylated RNaseB forms that were poorly recognized as substrates by GT were also less efficient inhibitors, whereas the insoluble derivatives that were better substrates were also better inhibitors of GT (Fig. 4 B).

Discussion
Reglucosylation of incompletely folded glycoproteins in the ER mediates their interaction with calnexin, calreticulin, and ERp57, which are part of a unique glycoproteinspecific chaperone system. Most glycoproteins are reglucosylated during their biogenesis (Gañan et al., 1991;Gotz et al., 1991), and several glycoproteins have been specifically shown to be reglucosylated in vivo during their maturation in the ER, including vesicular stomatitis G protein (Cannon and Helenuis, 1999;Suh et al., 1989), influenza HA (Hebert et al., 1995), transferrin (Wada et al., 1997), T cell receptor subunits (Van Leeuwen and Kearse, 1997), and cruzipain (Labriola et al., 1999). The reglucosylating enzyme uses only nonnative glycoproteins as substrates in vitro (Sousa et al., 1992;Trombetta and Parodi, 1992;Fernandez et al., 1994;Parker et al., 1995), recognizing occupied glycosylation sites covalently attached to nonnative protein backbones (Sousa et al., 1992;Trombetta and Parodi, 1992;Fernandez et al., 1994;Parker et al., 1995;Sousa and Parodi, 1995). However, little is known about the mechanism by which GT distinguishes between native and nonnative protein structures. To investigate the principles underlying the conformational sensor in the lectin-based chaperone system, we analyzed the interaction of GT with a series of defined conformers of a naturally occurring glycoprotein, RNaseB.
Intact native RNaseB or a form submitted to mock treatments, were neither substrates nor inhibitors of GT as expected. The nicked form, RNaseBS, was also not recognized despite having a partially perturbed conformation, judging by its distinct chromatographic behavior and its susceptibility to proteases and N -glycanase digestion. The increased exposure of the glycosylated Asn in an altered protein conformation was not sufficient for reglucosylation.
Removal of the NH 2 -terminal S-peptide (residues 1-20) from RNaseBS resulted, however, in a conformation that was recognized both as a substrate and inhibitor by GT. This conformer is an interesting and well-defined substrate. The soluble and monomeric nature of the RN-aseBS-Prot strongly indicates that some feature of its conformation is sensed as nonnative by GT. Like RNaseS-Prot (Richards and Wickoff, 1971;Richards et al., 1972;Shindo et al., 1979;Rosa and Richards, 1981), RNaseBS-Prot is likely to be highly structured. This is predicted by the constraints imposed by the four disulfide bonds and confirmed by its ability to rebind the S-peptide, which does not bind to denatured forms of RNaseBS-Prot (not shown). Although the affinity of the interaction is reduced, RNaseS-Prot is also known to bind RNase inhibitor (Neumann and Hofsteenge, 1994), an intimate association requiring proper RNaseA folding (Kobe and Deisenhofer, 1995). That GT specifically recognized the conformation of RNaseBS-Prot as nonnative, was shown by the stoichiometric correction of the conformation with the concomitant loss of the GT interaction by the rebinding of S-peptides. Therefore, the RNaseBS system reproduced the reglucosylation cycle observed in vivo using isolated components, underscoring the fact that it resembles the conformational requisites encountered in the lumen of the ER, where partially folded molecules become invisible to the enzyme upon proper folding.
Completely unfolded RNaseB molecules obtained by reduction and alkylation were poorly recognized by GT as substrates or inhibitors. That these were extensively denatured was shown by its enhanced susceptibility to proteases and N -glycanase, by their larger Stokes's radius observed by gel filtration, their ATP-dependent interaction with HSC70 (not shown), and their lack of ribonuclease activity. These results indicated that exposure of a glycosylated Asn, as in RNaseBS, even in combination with exposure of elements normally buried in native conformers, as in reduced and alkylated RNaseB, do not fulfill the requirements for recognition by GT.
Notably, we observed that GT interacted better with RNaseBS-Prot than with fully unfolded RNaseB. Some elements kept in close proximity in RNaseBS-Prot may become scattered in totally unfolded conformations, resulting in the observed loss of recognition by GT. Alternatively, GT may detect a structural scaffold in RNaseBS-Prot, resembling a partially structured or collapsed folding intermediate with limited flexibility, which would be largely absent in fully unfolded forms. If the loss of such structured conformation upon reduction of the disulfide bonds is what dampens recognition by GT, the enzyme could have a preference for structured folding intermediates as substrates. This possibility is indirectly supported by our observation that insoluble reduced/alkylated products of RNaseB interacted with GT better than the corresponding soluble forms. Also, in agreement with the observations presented here, using isolated components, reduced forms of glycosylated chimeras of bovine ribonuclease and bovine trypsin inhibitor are not reglucosylated when translated and translocated into microsomes in vitro (Rodan et al., 1996). Similarly, selective reglucosylation at advanced stages of folding after nearly complete oxidation-mediated binding of the trypanosomal protein cruzipain to calreticulin in vivo (Labriola et al., 1999).
Aside from the precise mechanism underlying the preference of GT for collapsed over fully unfolded conformations of RNaseB, we provide experimental evidence that GT not only distinguishes between native and nonnative conformations, but it can also discern between different nonnative conformations of the same glycoprotein. The finding that not all nonnative conformations of RNaseB are equally recognized by GT may be one of the reasons why protein unfolding, which was caused by the reduction of disulfide bonds, did not result in a generalized increase in protein reglucosylation in Schizosaccharomyces pombe (Fernandez et al., 1998).
Whereas chaperones of the Hsp70 family recognize short hydrophobic segments normally buried in native proteins (Zhu et al., 1996;Bukau and Horwich, 1998), our results suggest that GT prefers, instead, partially structured conformations. This observation correlates well with cases where reglucosylation was shown to mediate the participation of glycoproteins in the calnexin/calreticulin pathway. Influenza HA does not bind to BiP during its maturation, but is reglucosylated and binds calnexin and calreticulin (Hebert et al., 1995). On the other hand, vesicular stomatitis virus glycoprotein binds BiP during early stages of folding, and afterwards reglucosylation mediates its binding to calnexin (Hammond and Helenius, 1994). These examples show how the requirements for BiP binding and reglucosylation also differ in living cells. The development of defined substrates allows the design of novel experimental approaches to test these notions and to explore the mechanism underlying chaperone functions mediated by reglucosylation in the ER.