extfieldsofstudy: Biology Medicine

s2fieldsofstudy: Biology Medicine

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Oestradiol regulation of the components of the plasminogen-plasmin system in MDA-MB-231 human breast cancer cells stably expressing the oestrogen receptor.

To understand the hormonal regulation of the components of the plasminogen-plasmin system in human breast cancer, we examined the oestradiol (E2) regulation of plasminogen activators (PAs), namely urokinase-type plasminogen activator (uPA) and tissue-type plasminogen activator (tPA), plasminogen activator inhibitor type 1 (PAI-1) and uPA receptor (uPAR), in our model system. We used stable transfectants of the MDA-MB-231 human breast cancer cells that express either the wild-type (S30 cells) or the mutant 351asp-->tyr oestrogen receptor (ER) (BC-2 cells). Northern blot analysis showed that there was a concentration-dependent down-regulation of uPA, tPA and PAI-1 mRNAs by E2. In contrast, uPAR mRNA was not modulated by E2. The pure anti-oestrogen ICI 182,780 was able to block E2 action, indicating that the regulation of these genes is ER mediated. The E2 also inhibited the expression and secretion of uPA, tPA and PAI-1 proteins as determined by enzyme-linked immunosorbent assay (ELISA) in cell extracts (CEs) and conditioned media (CM). Zymography of the CM confirmed the inhibitory effect of E2 on uPA activity. Thus, we now report the regulation of uPA, PAI-1 and tPA by E2 in both mRNA and protein levels in ER transfectants. The association between down-regulation of the uPA by E2 and known E2-mediated growth inhibition of these cells was also explored. Our findings indicate that down-regulation of uPA by E2 is an upstream event of inhibitory effects of E2 on growth of these cells as the addition of exogenous uPA did not block the growth inhibition by E2. ImagesFigure 1Figure 2Figure 4Figure 6

tumour growth. invasion and metastasis is well documented (Kwaan. 1992. Tumour cell invasion is accomplished by the concerted action of several extracellular proteolytic enzyme systems. one of which is the plasminogenplasmin system. The different components of this system. e.g. urokinase-type plasminogen activator (uPA). its receptor (uPAR). tissue-type plasminogen activator (tPA) and the plasminogen activator inhibitor type 1 (PAI-1), along with other proteolytic enzymes. are involved in the process of activation of plasminogen to plasmin, which, directly or indirectly through the activation of other matrix metalloproteinases, degrade most components of the extracellular matrix and basement membrane.
The uPA/uPAR system plays a key role in tumour cell invasion and dissemination (Dan0 et al. 1985: Kwaan. 1992). Binding of both double chain uPA (tcuPA) and single-chain uPA (scuPA) to its receptor (uPAR) with the concomitant cell-surface binding of plasminogen enhances plasmin generation . uPAmediated proteolysis is modulated by PAI-1. the inhibitor for both uPA and tPA. PAI-I binding to receptor-bound uPA results in internalization of the uPA/PAI-l complexes (Olson et al. 1992 Whether the uPAR is intemalized at the same time (Bastholm et al. 1994) or the receptor just plays an enhancing role in internalization (Olson et al. 1992) is not clear.
In carcinoma of the breast, the level of uPA antigen. but not tPA antigen. in tumour homogenates was found to be a strong unfavourable prognostic factor for relapse and overall survival (Janicke et al, 1989(Janicke et al, : 1990Foekens et al. 1992;Sumiyoshi et al, 1992). High PAI-I and uPAR levels have also been found to be unfavourable (Duffy et al. 1988;Janicke et al. 1990;Foekens et al. 1992;Grondahl-Hansen et al, 1993. Several studies have found no correlation between high uPA (Grondahl-Hansen et al, 1997), PAI-l or uPAR levels and the oestrogen receptor (ER) status of the tumour (Duggan et al. 1995Gohring et al, 1995), whereas others (Foekens et al, 1994;Grondahl-Hansen et al. 1993;Ferno et al. 1996) have demonstrated that high uPA.
PAI-1 and uPAR contents in tumours are negatively correlated with the ER and progesterone receptor (PgR). In contrast, Duffy and colleagues (1986) state that levels of tPA activity had the highest correlation with ER and PgR positivity in human breast carcinomas. Determination of uPA levels and. to a lesser extent, PAI-1 levels was found to be useful in predicting the rate of response to tamoxifen therapy for metastatic disease . Cell lines derived from human carcinoma of breast have also been studied for the expression of uPA, uPAR. PAI-I and tPA (Butler et al. 1979;Shyamala and Dickerman. 1982;Huff andLippman. 1984: Mangel et al, 1988;Madsen andBriand. 1990: Holst-Hansen et al. 1996) and their role in cancer cell invasion and metastasis (Madsen and Briand, 1990;Holst-Hansen et al. 1996: Long and Oestadiol regulation of uPA, tPA, PAI-1 in breast cancer cells 89 Rose, 1996). The plasminogen activators' (PAs) activity in ERcontaining human breast cancer cell lines is stimulated by oestradiol (E,) and suppressed by anti-oestrogens (Butler et al, 1979;Shyamala and Dickerman, 1982;Huff and Lippman, 1984;Katzenellenbogen et al. 1984). Most investigators, with only one exception (Yang et al, 1983), have found that tPA and not uPA is regulated by E, in MCF-7 cells (Ryan et al, 1984;Dickerman et al, 1989;Butler et al, 1979;Mizoguchi et al, 1990). On the other hand, Mangel et al (1988) have shown that uPA was stimulated by E, in the T47-D and ZR-75-1 cells containing lower levels of ER, whereas MDA-MB-231 cells, which do not contain ER, showed a high level of both PAs activity that was not modulated by E,.
Interactions between oestrogen, tamoxifen and retinoic acid are also reflected in the expression of PAs by breast cancer cells (Butler and Fontana, 1992). PAI-1 levels were not influenced by E, in MCF-7 culture media (Davis et al, 1995), whereas in endometrial Ishikawa cells both E, and progesins induced the expression of PAI-1 and its mRNA (Fujimoto et al, 1996). Thus, in hormonedependent breast cancers, dissolution of the extracellular matrx may be modulated by PAs and PAI-1 under hormonal control.
To understand the hormonal regulation of components of plasminogen-plasmin system, we examined their E, regulation in our model system, using ER-negative MDA-MB-231 breast cancer cells transfected with either the wild-type (S30 cells) or the codon 351 AsT mutant ER (BC-2 cells). The purpose of this study was, using S30 and BC-2 cell lines, to: (a) examine the oestogen responsiveness of the components of the plasminogen-plasmin system, namely uPA, tPA, PAI-1 and uPAR; (b) determine if the process is ER-mediated; and (c) correlate the oestrogen responsiveness of the uPA with known growth inhibitory effect of E, on these cells.

Northem blot analysis
Northern blot analysis was performed as described previously (Levenson et al, 1997). Briefly, total RNA was isolated from cells following 48 h of treatment with compounds using the Trizol Reagent (Gibco, BRL). Twenty micrograms of total RNA sample was fractionated in 1.2% agarose-formaldehyde gel and transferred to a nylon membrane (Hybond-N+; Amersham, Arlington Heights, IL, USA). The membranes were hybridized at 420C with the corresponding 32P-labelled probes. The membranes were then washed and autoradiographed by exposure to Hyperfilm (Amersham) at -8C with intensifying screens. The expected 2.5-kb (uPA), 2.6-kb (tPA), 3.2-and 2.3-kb (PAI-1), and 1.4-kb (uPAR) tanscripts were detected. Because the sizes of mRNAs were similar, probing was performed on separate blots for each gene. Subsequently, the blots were stripped and reprobed with a cDNA to 1-actin. The signals were quantitated using phosphorimage analysis (Molecular Dynamics phosphorimager, Image Quant software).
Enzymelinked immunosorbent assay (ELISA) Two hundred thousand cells were plated per well in six-well dishes in 3 ml of media as described above. The cells were then treated with 10-9 M oestradiol or ethanol (control) for 48 h. Cell extracts (CEs) were isolated using Camiolo buffer (potassium acetate 75 mM, sodium chloride 300 mM, L-arginine 100 mM, EDTA 10 mM and 0.25% Triton X-100). The conditioned media (CM) were centrifuged at 5000 g and aliquots were stored at -00C until use. The corresponding cells were counted so that the levels of proteins could be standardized to the cell number. For the ELISA, the CM were diluted 1:10, and CEs were diluted 1:50 in PBS EDTAJTween and samples were analysed for uPA, tPA, PAI-l antigen levels using the ELISA kits TintElize uPA, tPA or PAI-I (Biopool, Sweden) with the corresponding antibodies. The uPA ELISA recognizes the scuPA as well as active tcuPA, whether it is free, receptor bound or complexed with PAI-1. For the quantitation of uPAR as well as uPAR/uPA and uPAR/uPA/PAI-l complexes the uPAR ELISA kit (American Diagnostica, Greenwich, CT, USA) was used. All assays were performed as described by the supplier.

Chromogenic assays
The proteolytic activity of uPA was assayed using a commercial Chromolize kit (Biopool, Ventura, CA, USA). The CM were added to a microtest plate well coated with monoclonal antibody against uPA, enabling the adsorption of uPA to the plate wall. After washing the non-absorbed material, plasmin was added to the well for the conversion of all scuPA to the active tcuPA. The uPA activity was determined by the addition of plasminogen and a plasmin-sensitive chromogenic substrate (D-But-CHT-Lys-pNA). uPA converted the plasminogen to plasmin, which reacts with the chromogenic substrate. This reaction was assayed by measurement of the absorbance at 405 um.
Zymography uPA, tPA and PAI-1/PA complex activities were analysed using zymography. Samples were electrphoresed in the presence of sodium dodecyl sulphate (SDS) under denaturing but non-reducing conditions in 10% polyacrylamide gels (SDS-PAGE) using the buffer system of Laemmli (1970) by a modification of the methods of Granrelli-Piperno and Reich (1978) and DePetro et al (1984).
Brffiish Journal of Cancer (1998) 78(1) uPA Diagnostica) were applied in each gel. In addition, standards for uPA/PAI-l complex and for tPA/PAI-l complex were applied. They were produced by first activating the 3.5 gg of PAI-I by boiling for 30 s, cooling slightly, adding excess uPA and tPA, respectively, and incubating at 37°C for 30 min to permit complex formation.
Growth assays Growth assays were performed as described previously (Levenson et al, 1997). Briefly, cells maintained in the media described above were plated at a density of 4x104 cells per well in 24-well plates and cultured for the 6 days either in media containing different concentrations of E,, uPA or in combinations. The media were changed every other day. Cells were then harvested and sonicated for 20 s with an ultrasonic cell disrupter.    Figure IA shows that expression of uPA mRNA is down-regulated by E, and that this effect is concentration dependent. In the presence of physiological concentrations of E, (IO9 M), there was a 3.6-fold decrease in uPA mRNA levels. At E, concentations of 106 M, the decrease was 7.4-fold. The expression of PAT-I mRNA was also down-regulated by E, in a concentration-dependent manner ( Figure IB). In the presence of 109 M E, there was a 1.7-fold decrease in PAI-I mRNA levels compared with unteated cells. In the case of tPA, the E, effect was even more pronounced ( Figure IC). In the presence of 1X9 M E,, there was 6.2-fold decrease compared with untreate cells. In contrast, uPAR mRNA renmained unaffected by E, at any given concentration ( Figure ID). To examine further the E,-mediated repression of uPA, tPA and PAI-I gene expression we used the pure antioestrogen ICI 182,780 alone and in combination with E, (Figure  2). The compound ICI 182,780 alone did not have any effect on these mRNA expressions, and in combination with E, was able to block E,-induced down-regulation of uPA, PAT-I and tPA mRNAs (Figure 2A-C). These data suggest that E, regulation of uPA, tPA and PAI-I genes occurs through the ER-mediated pathway. Once again, uPAR mRNA was not affected either by E, or by ICI 182,780 ( Figure 2D). The effect of E2 on evels ot uPA, PAM-1, tPA and uPAR as determned by EUSA CE and CM of S30 cels as descrbed Mateials and methods.
The data repesent dilleres between the l ev of comIponent in untreated contrl (100%) and levels in cels teated with 10-9 m E2 (% reduction) + s.e.m. *P< 0.005; *P= 0.06 in uPA levels in all cases. The uPA level when expressed as a ratio of E, treatment vs baseline was 0.49, indicating that E, has an inhibitory effect (Figure 3). This inhibitory effect of E, on uPA production reached statistical significance in CEs (P < 0.005) and less so in CM because of the variability of CM samples. Assay of PAI-I showed that E, also decreases levels of PAI-I in both CEs and CM, but to a lesser extent than uPA. Examination of tPA levels also revealed a definite decrease in this protease in expressed (CEs) and secreted (CM) levels in hormone-trea cells compared with controls ( Figure 3). By contrast, the uPAR levels in these cells were unaffected by E, treatment. We have previously shown that S30 and BC-2 cells are growth inhibited by 10-l-104 M concentrations of E, (Jiang and Jordan. 1992; Levenson et al, 1997). The fact that E, simultaneously inhibits endogenous uPA synthesis and activity while having no effect on uPAR prompted us to examine whether exogenous addition of intact uPA could (a) exert any effect on proliferative behaviour of S30 and BC-2 cells; and (b) reverse the inhibitory effect of E2 on growth. We performed a series of expenrments using various concentrations of uPA in a range from 10 nm to 5000 nm. The proliferative effect of added uPA was not different from that of untreated control cells at days 2, 4 and 6 (data not shown). In combination experiments with various concentrations of E,, uPA in 106 M (1000 nM) was not able to reverse the E,-inhibitory effect on cell growth ( Figure 5). To dissect possible reasons for added uPA failure, we performed both zymography ( Figure 6A) and ELISA ( Figure 6B) using CM from S30 cells treated with E,, uPA and combinations of E, and uPA from day 2 (Fig 6). E, inhibited activity of uPA. tPA and uPAJ PAI-1. and tPA/PAI-1 complexes. Interestingly. the addition of exogenous (lane 5) uPA alone resulted in its localization as both free uPA and as uPA/PAI-l complex. When added to E, (lane 6), the activity of uPA is decreased by both E2 and formation of the uPAIPAI-1 complex. The uPAIPAI-l complex is also decreased because of the inhibitory effect of E, on PAI-l levels. Samples from this experiment were examined using ELISA to determine the total (endogenous + exogenous) amount of uPA ( Figure 6B).

Effect of
Results showed that the addition of exogenous uPA incrased the total amount of uPA; however, when added to uPA. E,. was able to decrease amount of total uPA below control levels. high levels of uPA, PAI-I and uPAR were found to be independent prognostic factors with respect to relapse-free and/or overall survival, particularly in post-menopausal women (Gr6ndahl-Hansen et al, 1992;Spyratos et al, 1992;Gohring et al, 1995;Femo et al, 1996;Grondahl-Hansen et al, 1992). When analysed with respect to the ER status of the tumour, no clear answer was found: some found no correlation with the ER (Duggan et al, 1995;Foekens et al, 1995;Grondahl-Hansen et al, 1997) whereas others (Grondahl-Hansen et al, 1992;Foekens et al, 1994;Ferno et al, 1996) demonstrated negative correlation between high levels of prognostic factors and ER/PgR status of the tumour. As to the hormonal regulation of the components of the plasminogen-plasmin system in breast cancer cells in culture and its ER-mediated nature, the findings are controversial, especially for uPA. tPA but not uPA, PAI-1 or uPAR has been found to be regulated by E, in breast cancer cells containing the ER (Butler et al, 1979;Dickerman et al, 1989;Davis et al, 1995).
In the surgical specimens studied the expression of uPA in breast carcinoma was not in the tumour cells but in the myofibroblasts and other stromal cells (Nielsen et al, 1996). However, this picure is far from being clear. For example, xenografts of MDA-MB-231 cells in nude mice produced tmours that, in in situ hybridization studies, showed 'mRNA for human uPA in virtually all the cancer cells' (Romer et al, 1994). In either case, the stromal cells or the MDA-MB-231 cells do not express the ER. Thus, we wish to stress that we are evaluating a model system of uPA regulation in the laboratory that does not directly replicate the clinical situation in breast cancer. The availability of cells derived from MDA-MB-231 human breast cancer cells, which stably express ER and contain high levels of PAs, prompted us to examine the relationship between ER content and hormonal regulation of expression of the components of the plasminogen-plasmin system. To control at least three steps of E, regulation of final PA activity, e.g. from gene transcription to the enzyme activity, we concentrated on studying the regulation of mRNA accumulation, intra-and extracellular protein concentrations, and enzyme activities.
We herein report an observation that not only tPA but also uPA Because the PA protein levels do not necessarily reflect the PA activity, we performed zymographic analyses to examine the effect of E2 on the activity of uPA and tPA in the media conditioned by cells in the absence and pesence of 10-9 M E2. We found that E2 inhibits activity of uPA, tPA and PA/PAI-1 complexes. Thus, the inhibitory effect of E2 on PA activities in these cells occurs at any level examined from mRNA to enzyme activity.
It has been reported that uPAR is present in breast cancer tissue but not in normal breast tissue (Needham et al, 1987). uPAR is expressed in a variety of cancer cell lines and its synthesis is regulated by growth factors, such as epidennal growth factor (EGF), transforming-growth factors (TGF-1 and TGF-f2) and by the umour prmoter phorbol myristate acetate (Lumd et al, 1991a(Lumd et al, , 1995. In its native form uPAR is a glycolipid-anchored integral membrane protein. Therefore, we were interested in examining uPAR levels in cell extracts that represent potentially functional receptors compared with water-soluble tion products of uPAR found in cytosols (Gr6ndahl-Hansen et al, 1995). We found that our cells express uPAR, and that its synthesis is not rgulated by E2.
A mitogenic effect of uPA has been demonstra in vitro in different cell lines, both normal and neoplastic (Rabbai et al, 1990;He et aL 1991;De Petro et al, 1994;luperello et al, 1996).
Conversely, the inhibition of endogenously produced uPA by human malignant melanoma cells impairs cell proliferation (Kirchheimer et aL 1989). It is believed that the mitogenic and growth factor activity of uPA occurs throgh the amnotermnal fragment (AIT) of uPA, containing the growth factor domain, which shares sucntures homologous to the EGF, the TGF-a and the Krinkle domain. The ATF is a binding site of the uPA molecule for uPAR. The activity of uPA depends on the presence of uPAR on the cells as its mitogenic effect is selectively blocked by the addition of antibodies specific for the receptor (De Petro et al, 1994 this effect should be reversible upon the addition of exogenous uPA. However, the addition of exogenous uPA alone to culture media did not have any growth stimulatory effects on S30 and BC-2 cells, indicating that stimulatory effects of uPA varies with different cell types. When added to E2, uPA was not able to block the inhibitory effect of E2 on the growth. Tlere are several possible explanations: (a) down-regulation of uPA by E, is an upstream event of inhibitory effect of E2 on growth. Thus, replacement of uPA would not affect El-mediated growth inhibition. Parallel to our findings, Long et al showed (1996) that although EGF and TGF-a exerted stimulatory effects on uPA expression in S30 cells, neither of these growth factors was able to reverse the suppressive action of E, on growth of these cells. (b) Exogenously added uPA is inactivated by forming a uPA/PAI-l complex ( Figure  6A). It is possible that recombinant tcuPA added to the culture is more predisposed to complex formation with PAI-1 than endogenous cellular uPA, which is secreted as a scuPA. This can explain the detection of a large uPA/PAI-l band ( Figure 6A, lane 5) and subsequent absence of proliferative effect of uPA on these cells. Tlne addition of E, along with exogenous uPA to the culture affects endogenous uPA levels as well as uPA/PAI-l complex formation ( Figure 6A, lane 6).
In sumnmay, we have shown that uPA, tPA and PAI-1 are regulated by E, via the ER-mediated pathway in breast cancer cells stably transfected with the ER inreasing levels of endogenous uPA by transfection experiments with expression plasmids containing the uPA gene might be useful in future investigations.


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S-phase-specific activation of Cds1 kinase defines a subpathway of the checkpoint response in Schizosaccharomyces pombe.

Checkpoints that respond to DNA structure changes were originally defined by the inability of yeast mutants to prevent mitosis following DNA damage or S-phase arrest. Genetic analysis has subsequently identified subpathways of the DNA structure checkpoints, including the reversible arrest of DNA synthesis. Here, we show that the Cds1 kinase is required to slow S phase in the presence of DNA-damaging agents. Cds1 is phosphorylated and activated by S-phase arrest and activated by DNA damage during S phase, but not during G1 or G2. Activation of Cds1 during S phase is dependent on all six checkpoint Rad proteins, and Cds1 interacts both genetically and physically with Rad26. Unlike its Saccharomyces cerevisiae counterpart Rad53, Cds1 is not required for the mitotic arrest checkpoints and, thus, defines an S-phase specific subpathway of the checkpoint response. We propose a model for the DNA structure checkpoints that offers a new perspective on the function of the DNA structure checkpoint proteins. This model suggests that an intrinsic mechanism linking S phase and mitosis may function independently of the known checkpoint proteins.

Checkpoint pathways that respond to changes in DNA structure ensure the integrity of the DNA. After detection of specific DNA or DNA-protein structures, a signal is transduced to effector molecules that implement checkpoint-dependent responses such as cell-cycle arrest (Elledge 1996). Many components of the DNA-structure checkpoint pathways have been identified (Carr and Hoekstra 1995). In the fission yeast Schizosaccharomyces pombe, a group of six checkpoint Rad proteins (Rad1, Rad3, Rad9, Rad17, Rad26, and Hus1) are thought to participate in the monitoring and signaling processes that detect both DNA damage and incomplete DNA replication (Al-Khodairy and Carr 1992;Enoch et al. 1992;Rowley et al. 1992;Al-Khodairy et al. 1994). Central to this group is the Rad3 protein, which shares homology with both budding yeast and human checkpoint proteins (Savitsky et al. 1995;Bentley et al. 1996;Cimprich et al. 1996). Rad3 is a member of a larger subfamily of protein kinases that share structural similarities. This subfamily consists of large proteins with a lipid kinase-related domain at the carboxyl terminus. One member, DNA-P-Kcs, is well characterized as a protein kinase that is activated by association with DNA-binding subunits (Jeggo et al. 1995). By analogy with DNA-PK, we have proposed that Rad3 is activated by the other checkpoint Rad proteins, which may interact with the specific DNA or DNA-protein structures generated by DNA damage and DNA synthesis (Carr 1997).
DNA structure checkpoints respond to several distinct signals. The best characterized are DNA damage caused by UV or ␥-irradiation and S-phase arrest resulting from hydroxyurea (HU) exposure. In response to DNA damage, but not S-phase arrest, Chk1 kinase becomes phosphorylated in a manner dependent on Rad3 and the other checkpoint Rad proteins (Walworth and Bernards 1996). Thus, in this case, an effector is activated only in response to specific inputs. Thus, the damage and replication checkpoints can be formally considered as distinct pathways.
The DNA structure checkpoints were first identified by their effects on mitosis (Weinert and Hartwell 1988;Enoch and Nurse 1990). The checkpoint pathways, however, control other responses. For example, following treatment with HU, the checkpoint Rad proteins are essential to enable cells to survive S-phase arrest. In the absence of the checkpoint Rad proteins, HU causes rapid cell death during S phase (Enoch et al. 1992;Carr 1994), even though DNA synthesis can continue after the HU is removed. Thus, checkpoint-dependent reversible Sphase arrest is vital to survival when replication is perturbed. A related phenotype is also seen when checkpoint mutants are exposed to DNA damage during S phase: Survival of S-phase cells is dependent on a functional checkpoint pathway (Al-Khodairy et al. 1994).
In Saccharomyces cerevisiae, two proteins (Mec1 and Rad53) have been identified that are involved in both the DNA damage and the DNA replication checkpoints. Mec1 (Kato and Ogawa 1994;Weinert et al. 1994) is a homolog of Schizosaccharomyces pombe Rad3. Rad53 is a protein kinase that is phosphorylated and activated in response to DNA damage and HU exposure (Allen et al. 1994;Sanchez et al. 1996;Sun et al. 1996). Rad53 activation is dependent on Mec1, but null mutants of RAD53 are only partially defective in G 2 arrest after DNA damage (Pati et al. 1997). In S. pombe, a structural homolog of Rad53, called Cds1, has been identified as a multicopy suppresser of a temperature-sensitive DNA polymerase ␣ mutation (Murakami and Okayama 1995). Cds1 was proposed to link DNA replication to the mitotic machinery. Cds1 is not required to prevent mitosis following DNA damage.
In this report we show that Cds1 is a protein kinase that is phosphorylated and activated by DNA-replication arrest and activated by DNA damage. The activation by DNA damage is only seen in S phase, suggesting that the Cds1 response is specific to replication structures. The major effect of Cds1 deletion is not the inappropriate onset of mitosis, but an inability of cells to properly regulate S phase when challenged during replication. We propose a model for Cds1 function in which Cds1 defines a subpathway of the DNA structure checkpoints that prevents irreversible lethal DNA damage from occurring when DNA synthesis is perturbed.

S-phase arrest causes mitotic delay in the absence of Cds1
Cds1 has been reported to link S phase to mitosis (Murakami and Okayama 1995). In cds1 null mutants arrested in S phase by HU, however, mitotic arrest is essentially normal during the first 6 hr ( Fig. 1A and Fig. 3 of Murakami and Okayama 1995), approximately the length of time that HU delays bulk DNA synthesis (Sazer and Sherwood 1990). Despite this mitotic delay, there is rapid and irreversible loss of viability associated with entry into S phase in the presence of HU (Murakami and Okayama 1995). Rapid loss of viability during S phase is also seen with the checkpoint rad class of mutants (Enoch et al. 1992;Carr 1994).

Cds1 is required for the response to DNA damage
It has been reported that cds1 mutant cells are not sen-sitive to DNA damage (Murakami and Okayama 1995). These studies used UV as the DNA-damaging agent. We find that Cds1 null cells are significantly sensitive to UV (Fig. 1B) and ionizing radiation (Fig. 1C), despite the fact that they maintain a normal G 2 checkpoint when irradiated (Fig. 1D). To characterize this sensitivity further, we have created double mutants with a number of well characterized checkpoint mutants (Fig. 1B,C).
The Chk1 mutant is unable to arrest mitosis after DNA damage, but is more resistant to radiation than the checkpoint rad group of mutants (Al-Khodairy et al. 1994). Interestingly, a specific allele of one of the checkpoint rad genes, rad26.T12, has been characterized that retains a normal mitotic arrest after DNA damage, but is still sensitive to DNA-damaging agents (Al-Khodairy et al. 1994). Genetic studies of chk1.d rad26.T12 double mutants show that these cells have a similar phenotype to the rad26.d null mutant. This led us to propose that rad26.T12 is defective in a subpathway of the checkpoint response that is independent of mitotic arrest (Al- Khodairy et al. 1994).
We have created cds1.d chk1.d double mutants and compared the sensitivities with the respective single mutants and with the rad26.d null mutant. cds1.d chk1.d double-mutant cells are more sensitive to DNA damage than the two single mutants, as would be expected from their distinct phenotypes. The double mutant shows an equivalent sensitivity to rad26.d (Fig. 1B,C). We have also created cds1.d rad26.T12 double mutants and compared the sensitivity of these with the respective single mutants. The double mutants are no more sensitive to DNA damage than the most sensitive single mutant (Fig. 1B,C). This is also true for the cds1.d rad26.d double mutant (data not shown).
The classical interpretation of epistasis analysis in repair studies is as follows: Two mutants that do not give additional sensitivity when combined (compared with the most sensitive single mutant) act in the same pathway. Conversely, two null mutations that give increased sensitivity when combined act in distinct pathways. The simplest interpretation of our data is that the cds1-dependent DNA-damage response and the chk1-dependent DNA-damage response both require the correct function of the checkpoint rad pathway. Therefore, we can separate the checkpoint rad pathway into two subpathways, one mediated by Chk1, which arrests mitosis, and one mediated by Cds1, which is independent of mitotic arrest. rad26.T12 mutant cells would appear to be defective in the Cds1, but not the Chk1, dependent pathway. We will refer to the Cds1-mediated response as the checkpoint rad-dependent S-phase recovery pathway.
cds1 is a multicopy suppressor of rad26.T12 Sequence analysis of the rad26.T12 allele identified a single arginine to tryptophan mutation at position 541 of the Rad26 protein (see Materials and Methods). This site is 74 amino acids from the carboxyl terminus of the protein. An allele of rad26 with a defect in sensing slowed DNA replication has been described recently (Uchiyama et al. 1997). This phenotype results from a duplication of 6 amino acids within Rad26, at position 511, 30 amino acids distant from the rad26.T12 mutation. To investigate the potential involvement of a carboxy-terminal Rad26 domain in the checkpoint rad-dependent recovery pathway, we deleted incremental carboxy-terminal regions. Removal of the carboxy-terminal 11 amino acids of rad26 (604-614) did not affect function. Removal of the 31 amino acids (584-614) resulted in a phenotype similar to the null mutant ( Fig. 2B; see Materials and Methods), despite the fact that a correct size protein is produced (data not shown). Therefore, the carboxyl terminus does not encode a defined functional domain involved in the recovery pathway.
To further investigate the relationship between Cds1 and the checkpoint rad-dependent recovery pathway, we transformed rad26.d and rad26.T12 cells with a plasmid containing a cds1 genomic clone. As controls, we used an empty vector and a wild-type rad26 gene. cds1 overexpression rescues much of the radiation sensitivity and HU sensitivity of the rad26.T12 allele ( Fig. 2A), consistent with Cds1 functioning either in parallel or downstream of Rad26. This effect is unlikely to be the result of any cell-cycle delay imposed by the overexpression, as cell-cycle delay following irradiation is proficient in the rad26.T12 strain (Al-Khodairy et al. 1994). Furthermore, Cds1 does not unduly affect the cell cycle as judged by cell length at mitosis in these experiments.
The common involvement of Rad26 and Cds1 in the   2C). HA-Rad26 is able to precipitate Myc-Cds1, and HA-Cds1 is able to precipitate Myc-Rad26. In addition, Figure 2. cds1 is a multicopy suppresser of rad26.T12 mutant cells. (A) rad26 null and rad26.T12 mutant cells were transformed with rad26 genomic DNA, cds1 genomic DNA, or an empty vector control (pAL). Cells were serially diluted (from 10 7 cells/ml to 10 4 cells/ml), spotted onto plates and irradiated at the doses indicated. Cells were also spotted onto plates containing a range of HU concentrations. (B) rad26 null cells were transformed with the rad26 genomic clone, the empty vector (pAL) or rad26 constructs containing carboxy-terminal deletions. Numbers correspond to the length of the constructs (amino acids). Full-length Rad26 is 614 amino acids. Cells were diluted to the concentrations shown and either spotted onto plates and irradiated at the doses indicated or spotted onto plates containing different concentrations of HU. (C) Cds1 immunoprecipitates with Rad26 in S. pombe extracts. Wild-type S. pombe was transformed with REP41 (leucine selection) and REP42 (uracil selection) plasmids containing cds1 and rad26 cDNA tagged with either HA or Myc epitopes. Immunoprecipitation was carried out on extracts from logarithmically growing cells, after 18 hr growth without thiamine. Immunocomplexes were processed for Western blotting with anti-Myc antibody. Rad26 associates with itself (lane 3) and with Cds1 (lanes 7,8). The empty vector (lanes 1,2,4,5) was used as control. We estimate overexpression at ∼50× of wild-type protein levels. Comparing the amount of Cds1 precipitated with Rad26 to total protein, we estimate ∼3%-5% recovery. (D) Immunoprecipitations were repeated by use of proteins expressed in rabbit reticulocyte lysate. In this case, Rad26 was seen to associate with itself, but not with Cds1, suggesting bridging proteins or posttranslational modification are required for the Rad26-Cds1 interaction. This would be consistent with levels of recovery.
Rad26, but not Cds1, appears to form homomultimers, as HA-Rad26 precipitates Myc-Rad26 when the two constructs are expressed in the same cell. To determine if these interactions were direct, or require either bridging proteins or specific posttranslational modification, we translated Rad26 and Cds1 in rabbit reticulocyte lysates for coimmunoprecipitation (Fig. 2D). Rad26 did not precipitate with Cds1, but still interacted with itself. Thus, we conclude that bridging proteins or modifications are required for Cds1-Rad26, but not Rad26-Rad26 interactions.

Cds1 is a protein kinase and phosphoprotein activated by S-phase arrest and DNA damage
To verify that Cds1 is a protein kinase, we purified overexpressed GST-Cds1 fusion protein from S. pombe and established conditions under which it will phosphorylate myelin basic protein (MBP) in vitro (data not shown). Specificity for Cds1 was shown by an equivalent purification of a GST fusion protein carrying an Asp to Glu mutation in residue 312 of the kinase domain. This kinase dead protein did not have kinase activity for MBP (data not shown). A rabbit polyclonal antibody to Cds1 protein was used to detect Cds1 protein in asynchronous wild-type cells. By immunoprecipitation (IP) and Western blotting after SDS-PAGE, a single band is seen that migrates at ∼50 kD, near the predicted molecular weight of 51.9 kD for Cds1 (Fig. 3A). This band is absent in IP's from cds1 null mutant cells. In addition, the antibody recognizes overexpressed Cds1 (data not shown), verifying its specificity. Following incubation of cells in HU, a mobility shift for Cds1 is evident by SDS-PAGE; Cds1 migrates as a more diffuse band, with an apparent increase in molecular mass of several kilodaltons (Fig. 3A,B). The mobility shift can be reversed by phosphatase treatment (Fig. 3B), indicating that it is a result of phosphorylation. To ascertain if Cds1 phosphorylation correlates with increased kinase activity, IP kinase assays against an MBP substrate was used to measure the Cds1-dependent kinase activity from treated and untreated cells. Cds1-dependent activity increases markedly in IP kinase assays performed on extracts prepared from cells treated with HU ( Fig. 3A). By overexpressing Cds1 and kinase-dead Cds1 ∼25-fold in cds1 null cells, we have shown that the activity observed is dependent on the integrity of the Cds1 kinase domain (Fig. 3A). These data indicate that and kinase activity (bottom) from wild-type and Cds1 null cells by use of affinity-purified Cds1 sera or preimmune sera identifies Cds1 as a single band migrating at 52 kD. In extracts from cells treated with HU, Cds1 undergoes an apparent increase in molecular weight (second lane). Kinase activity against MBP in the IPs correlates with the presence of Cds1 and increases in cells treated with HU. To show the specificity of the kinase activity to Cds1, wild-type or kinase-dead Cds1 protein were expressed in cds1 null cells, immunoprecipitated (top, right) and assayed for kinase activity (bottom, right). (B) The Cds1 mobility shift can be reversed by phosphatase treatment. Ten and twenty units of CIAP restored the mobility of Cds1 from HU-treated cells to that of Cds1 from untreated cells. Twenty units of heat-treated CIAP does not affect Cds1 mobility. (C) Cds1 kinase can be activated by DNA damage. Asynchronous cultures of wild-type and chk1 null cells were subjected to 500 Gy of ionizing radiation. Cell extracts were made at time points following irradiation and assayed for Cds1 kinase activity. Wild-type cells showed a small but reproducible increase in Cds1 kinase activity. chk1 cells showed a greater increase in Cds1 kinase activity. (D,E) Cds1 phosphorylation (D) and activation (E) following HU treatment is dependent on the function of the checkpoint Rad proteins. Cds1 phosphorylation is not seen in immunoprecipitates from rad1, rad3, rad9, rad17, rad26, or hus1 null mutants but is evident in wild-type (WT) and chk1 mutants. IP-kinase activity is much reduced or absent in the checkpoint rad null strains when compared with wild-type and chk1 null cells. the phosphorylation event signifies the activation of the Cds1 kinase.
Because cds1.d mutants are sensitive to DNA damage, we have investigated Cds1 activation following treatment with ionizing radiation. By use of IP kinase assays, we can detect activation of the Cds1 kinase in wild-type cells after irradiation (Fig. 3C). Because wild-type cells accumulate in G 2 following irradiation, we have also tested chk1 mutant cells, which do not arrest in G 2 after DNA damage (Al-Khodairy et al. 1994). Increased levels of activation were seen. We have been unable to detect a mobility shift of Cds1 following DNA damage (data not shown). The kinase activity of Cds1 following DNA damage does not reach the levels seen following treatment with HU. Thus, we cannot determine if the phosphorylation event observed following HU treatment is distinct from that which occurs after DNA damage, or if Cds1 is partially phosphorylated in response to DNA damage.

The increases in Cds1 phosphorylation and kinase activity are dependent on the checkpoint Rad proteins
The Cds1 homolog in S. cerevisiae, Rad53, is a phosphoprotein that is activated by HU treatment and DNA damage. Rad53 activation is dependent on Mec1, the homolog of S. pombe Rad3 (Sanchez et al. 1996;Sun et al. 1996). By use of null mutants of the six checkpoint rad genes, we have shown that the phosphorylation (Fig. 3D) and activation (Fig. 3E) of Cds1 induced by HU is largely dependent on the function of the six checkpoint Rad proteins, Rad1, Rad3, Rad9, Rad17, Rad26, and Hus1. Both rad3.d and rad26.d mutants are completely unable to phosphorylate or activate Cds1. Low, but reproducible, levels of activation are seen in rad9.d, rad17.d, and hus1.d mutants. A moderate level of activity is seen in rad1.d cells. In no case was a Cds1 mobility shift seen.
Our genetic analysis places Cds1 function in the checkpoint rad-dependent recovery pathway, a subpathway of the checkpoint response. This subpathway has also been identified as defective in the rad26.T12 mutant, and we find evidence of an association between Cds1 and Rad26. It is therefore possible that the major function of Rad26 is in the recovery pathway. To address this, we followed the kinetics of Cds1 phosphorylation and activation in response to HU in rad26.T12 cells (Fig.  4A). Compared with wild type, there is a reduction in Cds1 activity, and Cds1 does not become phosphorylated. Quantification (Fig. 4B) of Cds1 kinase activity shows less than half the activity seen in wild-type cells. These data are consistent with a role for Rad26 in mediating the Cds1 response.

Cds1 is not active during normal S phase
In S. cerevisiae, it is not clear if Rad53 is activated during S phase of the normal cell cycle (Sanchez et al. 1996;Sun et al. 1996). If Cds1 is activated and phosphorylated during unperturbed S phase, the increased activity following HU treatment or DNA damage may be a consequence of cells accumulating in S phase. We have studied Cds1 phosphorylation and activity in synchronized cells. Cells were synchronized in two ways; either by a temperature shift and release strategy with a temperature-sensitive cdc10.V50 mutant [cdc10 mutants arrest the cell cycle at start, during G 1 (Aves et al. 1985)] or by centrifugal elutriation. No evidence of either a mobility shift or reproducible activation of Cds1 kinase was seen as cells proceeded through unperturbed S phase (Fig. 5B,D; data not shown). On occasion, low levels of Cds1 activity, but not phosphorylation, are detected in late S phase [i.e., Fig. 5B, 90(−)]. This is not consistent, however, and may represent experimental artefacts or low levels of spontaneous replication stalling in synchronous cultures.
The most likely explanation for Cds1 activation during HU treatment is that Cds1 is activated in response to stalled DNA replication. To rule out the alternative possibility that HU treatment itself will activate Cds1 irrespective of cell-cycle position, we synchronized wildtype cells by elutriation and treated aliquots with HU during either G 2 or G 1 /S phase. We observed that the activation of Cds1 and its associated mobility shift (data not shown) occurs at a time commensurate with the effect of HU during S phase (Fig. 5A,B), and is not an effect of the HU itself. Thus, we conclude that HU induced Cds1 activation is specific to perturbed S phase.

Cds1 is activated by DNA damage during S phase
In S. cerevisiae, Rad53 is activated by DNA damage during G 1 , S phase, and post S phase. Because there are significant phenotypic differences between rad53 and cds1 mutants in the two yeasts, we investigated the cell-cycle specificity of Cds1 activation by DNA damage. In synchronous wild-type cells generated by elutriation, Cds1 activation was only seen when cells were irradiated during the G 1 /S period of the cell cycle, and did not occur in cells irradiated during G 2 phase (Fig. 5C,D at 0 min). Thus, Cds1 activation occurs only in G 1 and/or S phase. To distinguish between G 1 and S phase, we arrested cells at start with the cdc10.V50 mutant before treating them with radiation (Fig. 5E). G 1 arrested cells did not activate Cds1 after irradiation when held in G 1 . These two experiments allow us to conclude that Cds1 is activated by DNA damage only during S phase.

Cds1 activation in a range of temperature-sensitive cdc mutants
To define more precisely at which point during S phase Cds1 is able to be activated, we have studied the activation of Cds1 in a range of temperature-sensitive cdc mutants defective in various replication proteins (Fig. 5F). We find that Cds1 is strongly activated only in temperature-sensitive polymerase ␣ (D' Urso et al. 1995;Francesconi et al. 1997) and temperature-sensitive polymerase ⑀ (cdc20) mutants following a temperature shift. Cds1 is not activated in temperature-sensitive polymerase ␦ (cdc6) (Iino and Yamamoto 1997) or in cdc21 (Mcm protein) (Mairorano et al. 1996) and is only moderately activated in late S phase cdc17 (DNA ligase) mutant cells (Barker et al. 1987). Cds1 is activated in cdc22 (ribonucleotide reductase) mutants (Fernandez-Sarabia et al. 1993), consistent with the data by use of HU.
Cds1 null mutants are defective in an S phase DNA-damage checkpoint S. cerevisiae mec1 and rad53 mutants are defective in slowing down S phase when treated with the DNA-dam- Figure 5. Activation of Cds1 kinase is S-phase dependent. (A) Septation of wildtype cells synchronized in G 2 by elutriation. The culture was divided into three and incubated at 30°. HU was added (20 mM) at either t = 0 (ᮀ) or t = 60 (᭹) min; control (). The cell-cycle stage of the culture was followed by septation index by Calcofluor and DAPI staining. Untreated cells show two peaks in septation. The second peak is missing in the HU-treated culture indicating the activation of the replication checkpoint. (B) Samples were taken from all three cultures and processed for Cds1 kinase assay. Cds1 kinase does not activate during G 2 despite the presence of 20 mM HU (30+, 60+). Cds1 kinase activity accumulates after 90 min when the majority of the cells have entered or are entering S phase. (C) Septation index of wild-type cells synchronized in G 2 by elutriation (30°C). (D) Cds1 kinase activity of irradiated or unirradiated cells removed at the times indicated and treated with 250 Gy of radiation. Cell extracts were prepared after 40 min of recovery incubation at 30°C. Cells irradiated in G 2 at t = 0 do not activate Cds1 kinase after irradiation, whereas cells entering S phase after irradiation develop significant activity. Consistent with the S-phase dependency of the kinase activation cells irradiated late in S phase/G 2 (t = 90,110) fail to generate significant kinase activity compared with the unirradiated control. (E) cdc10.V50 cells irradiated at the permissive temperature (26°C) show activation of Cds1 kinase whereas cells irradiated and held at the restrictive temperature (37°C) do not, thus showing that Cds1 kinase is not activated by DNA damage during G 1 phase. (F) Activation of Cds1 by temperature shift in various cdc mutants defective in DNA synthesis. Logarithmic cultures of wild-type, cdc6, cdc17, cdc20, cdc21, cdc22, and pol␣ cells grown at 27°C were split into two samples, one was incubated for a further 2 hr, 30 min at 27°C, whereas the other was incubated at 37°C for 2 hr, 30 min before extracts were prepared and tested for IP kinase activity. aging agent methylmethane sulfonate (MMS). We have tested the ability of wild-type, rad3, chk1, cds1, rad26, and rad26.T12 mutant cells to slow S phase in response to MMS. By use of two different methods to synchronize cells in G 1 , we have shown that wild-type cells have this checkpoint response, and that it is lost in rad3 mutants (Fig. 6A,B). By use of one of the synchronization methods, we have further shown that cds1 mutant cells, but not chk1 mutant cells, have also lost this checkpoint. As expected, rad26 null mutant cells behave in a similar manner to rad3. Consistent with our genetic data, rad26.T12 mutant cells are also defective in slowing S phase when DNA is damaged (Fig. 6B). These data are consistent with Cds1 and Chk1 defining specific sub-pathways of the checkpoint Rad response. They also show that rad26.T12 mutants are specifically defective in the Cds1-dependent, but not the Chk1-dependent response.

Loss of Cds1 results in the abnormal activation of Chk1 by HU
A clue to the biological significance of Cds1 activation can be seen by examining the phenotype of the cds1.d chk1.d double mutant, and the behavior of the DNA damage-specific Chk1 kinase in cds1 null cells. Unexpectedly, after HU treatment, cds1.d chk1.d double mutants enter mitotic catastrophe and form cuts in a man- Figure 6. The S-phase progression checkpoint requires rad3, rad26, and cds1. (A) rad + cells (h − cyr1 sxa2) and rad3 mutant cells (h − cyr1 sxa2 rad3) were synchronized in G 1 by centrifugal elutriation (collecting early G 2 cells) followed by exposure to P factor. Cells were released from P factorinduced G 1 arrest in either the presence or absence of 0.033% MMS and S-phase progression followed by FACS. Attempts to synchronize other checkpoint mutants this way were complicated by the morphological changes associated with the cyr1 sxa2 background. (B) Wild-type, rad3, cds1, chk1, rad26, and rad26-T12 cells were synchronized in G 1 by nitrogen starvation, released into the cell cycle in the presence or absence of 0.033% MMS and followed by FACS. As before, rad3 cells are unable to slow S phase in response to DNA damage. Similarly, cds1 and rad26 are required for this checkpoint, whereas chk1 is not. Cells carrying the rad26.T12 mutation also have a defect in this checkpoint. ner identical to the checkpoint rad null mutants such as rad3.d and rad26.d (Fig. 7A). From this analysis, it is apparent that the mitotic delay seen in cds1 null mutants is dependent on Chk1, which is primarily involved in the DNA-damage checkpoint at mitosis (Walworth et al. 1993;Al-Khodairy et al. 1994). There are two possible explanations for this. Either Cds1 and Chk1 play redundant overlapping roles to arrest mitosis in the presence of the S-phase inhibitor HU, or the Chk1 kinase is activated when cds1 null mutant cells are treated with HU.
Chk1 kinase has been shown to be phosphorylated in response to DNA damage, and this phosphorylation event correlates to Chk1 function. Furthermore, Chk1 has been shown not to be phosphorylated in response to HU treatment, which is consistent with its lack of a HU-sensitive phenotype (Walworth and Bernards 1996). These observations allow us to distinguish between the two possibilities discussed above. We have created a cds1 null mutant strain, in which the chk1 gene is tagged in the genome by three HA epitopes (Walworth and Bernards 1996) and compared the phosphorylation of Chk1 in the presence and absence of Cds1 function following HU treatment (Fig. 7B). Chk1 becomes rapidly phosphorylated in response to HU only when Cds1 is absent, and not when it is present. Thus, the function of Cds1 during HU treatment prevents a situation in which Chk1 becomes phosphorylated. Because Chk1 normally responds specifically to DNA damage, we speculate that the activity of Cds1 prevents the occurrence of DNAdamage structures during S-phase arrest.

Discussion
We have shown that HU and DNA damage both activate the Cds1 kinase during S phase, but not during G 1 or G 2 . At least in the case of HU arrest, this is reflected by a phosphorylation event that results in a mobility shift of Cds1 on SDS-PAGE. We have also shown that Cds1 is activated at the restrictive temperature in temperaturesensitive mutants of DNA polymerase ␣ and ⑀, but not DNA polymerase ␦. Cds1 activation is dependent on the correct functioning of the checkpoint rad pathway, and Cds1 is able to interact with Rad26. Whereas cds1 mutant cells have a normal mitotic checkpoint after DNA damage, they are defective in regulating S phase in the presence of DNA-damaging agents. Together, these data show that the cds1 defines a subpathway of the checkpoint rad response that is specific to S phase. The evolutionary conservation of the DNA structure-dependent checkpoint pathways may have significant consequences for our understanding of carcinogenesis. During evolution there have been significant alterations in checkpoint protein function between different species (Carr and Hoekstra 1995). This is most clearly documented for the large lipid kinase motif proteins, such as Rad3 (S. pombe), Mec1/Tel1 (S. cerevisiae), and Atm/Atr (mammalian species) (Zakian 1995). Our data suggest that the same is true for the Cds1-Rad53 kinases.

Activation of Cds1 is specific to S phase
In S. cerevisiae, Rad53 is involved in all the known DNA structure checkpoint responses including G 1 and G 2 arrest after DNA damage, and G 2 arrest during HU treatment (Allen et al. 1994). The Rad53 protein can be phosphorylated and activated in response to DNA damage at different points in the cell cycle, and is similarly phosphorylated and activated during S phase and G 2 arrest (Sanchez et al. 1996;Sun et al. 1996). In contrast, Cds1 is not involved in the DNA damage-dependent arrest of mitosis and is only activated by DNA damage during S phase. Furthermore, whereas S-phase arrest by HU or The phosphorylation-dependent mobility shift of Chk1 kinase was assayed in cds1 + and cds1 null mutant cells containing an integrated triple HA-tagged chk1 gene (Walworth and Bernards 1996). Samples from asynchronous cultures treated with 12 mM HU were taken at the times indicated. Chk1 was detected, as described previously (Walworth and Bernards 1996), with anti-HA monoclonal. In cds1 + cells, Chk1 remains unphosphorylated, whereas in cds1 null mutant cells Chk1 is rapidly phosphorylated. temperature-sensitive polymerase ␣ and ⑀ mutants causes activation of Cds1, the cds1 null mutant does not have a defect in mitotic arrest when S phase is blocked by HU. Interestingly, a temperature-sensitive DNA polymerase ␦ mutant did not result in Cds1 activation at the restrictive temperature, and activation was low in mutants arrested in late S phase.

Cds1 defines a subpathway of the DNA-structure checkpoint response
The phosphorylation and activation of Rad53 in response to S-phase arrest is dependent on Mec1, and in response to DNA damage is dependent on Mec1 and Mec3 (Sanchez et al. 1996;Sun et al. 1996). Furthermore, Tel1 overexpression can restore HU and damage dependent Rad53 phosphorylation (Sanchez et al. 1996). In S. pombe, we find that Cds1 activation and phosphorylation in response to HU arrest is dependent on all six of the checkpoint Rad proteins. In S. pombe, Rad1 and Rad17 are required for all the checkpoint-dependent responses to DNA damage and S-phase arrest (Al-Khodairy and Carr 1992), whereas in S. cerevisiae the homologous proteins are only required for the DNA-damage responses and not the HU response (Weinert et al. 1994). Interestingly, a slight activation of Cds1 is evident in rad1, rad9, rad17, and hus1 null mutants.
Genetic analysis shows that Cds1 acts in a separate branch of the checkpoint response from Chk1. Combining the phenotypic observations of chk1 and cds1 mutants with the biochemistry of the kinases suggests that the checkpoint Rad proteins are involved in sensing and signaling the presence of specific DNA or DNA protein structures, and activate Chk1 or Cds1 as appropriate. Chk1 appears to be activated by DNA damage-specific structures, whereas Cds1 is activated by DNA replication-specific structure. This could reflect cell cycle and DNA metabolism-specific localization of the checkpoint Rad proteins, and suggests that the putative complex(es) formed between these proteins are capable of monitoring a range of distinct structures. By comparison with DNA-PK, in which the catalytic subunit (also a large lipid kinase motif protein) is activated by the Ku heterodimer when it associates with DNA ends, it is interesting to speculate that the Rad3 kinase is activated by association with distinct subunits (perhaps Rad1, Rad9, Rad17, Rad26, and Hus1), which may interact with different DNA or DNA-protein structures. In this context, it would also be possible to envisage an explanation for the different requirements in S. pombe and S. cerevisiae for these proteins. Perhaps in S. pombe, Rad3 function is compromised by loss of any associated subunits, whereas in S. cerevisiae, Mec1 function could be more tolerant.
The genetics and cell-cycle specific activation of Cds1 offers an opportunity to dissect out the specific function of Cds1 within the network of closely related checkpoint responses. Cds1 acts primarily to ensure the reversible arrest of DNA synthesis. This subpathway of the checkpoint rad response was first observed during studies on the hus1 mutant (Enoch et al. 1992). hus1 null mutants, in common with other checkpoint rad mutants, lose viability very rapidly on exposure to HU during S phase. Furthermore, by use of the rad26.T12 mutant, which retains a normal G 2 arrest after DNA damage (Al-Khodairy et al. 1994), it has been possible to associate an element of the radiation sensitivity seen in checkpoint rad mutants with radiation exposure during the G 1 /S phase of the cell cycle. Because epistasis and biochemical analysis shows that the radiation sensitivity associated with loss of Cds1 function is caused by a defect in the same pathway as that defined by the rad26.T12 mutation, and because cds1 and rad26.T12 mutants share a defect in slowing S phase when exposed to DNA-damaging agents, it is now possible to formally propose two distinct checkpoint rad dependent DNA metabolism responses: Chk1 mediated G 2 arrest and Cds1 mediated S-phase recovery.

Cds1 function may prevent DNA damage accumulating when S phase is arrested
Contrary to the published report, we show that cds1 null mutant cells do not have a significant defect in mitotic arrest following HU exposure. cds1 cells, however, die during S-phase arrest, in a manner independent of mitosis (Murakami and Okayama 1995). A further effect of HU treatment in cds1 null mutant cells is the activation of the DNA-damage checkpoint kinase Chk1. Because the double mutant cds1.d chk1.d does not arrest mitosis after exposure to HU, we can conclude that this unscheduled activation of Chk1 in response to HU is the reason cds1 single-mutant cells arrest mitosis in response to HU. The fact that Chk1 is not normally phosphorylated when cells are treated with HU, indicates that this is not because Cds1 and Chk1 kinase play overlapping redundant roles during S-phase arrest. Instead, we propose that the inappropriate activation of the Chk1 kinase in cds1.d cells in response to HU is a consequence of the accumulation of unrepairable DNA-damage structures during S-phase arrest, which, in turn, activate the DNA-damage checkpoint.

Cds1 may interact with replication proteins to inhibit the formation of, or stabilize, replication structures during S-phase perturbations
Cds1 acts downstream of the checkpoint Rad group of proteins and enables cells to arrest DNA synthesis reversibly in response to HU or DNA damage, thereby preventing the accumulation of unrepairable DNA lesions. It is not yet clear what the substrates of Cds1 kinase are. Possible substrates would be the components of the replication apparatus such as polymerase ␣-primase (Marini et al. 1997) or even upstream activators of replicon initiation such as the Hsk1 kinase (the S. pombe homolog of S. cerevisiae Cdc7) (Masai et al. 1995). In response to DNA-replication inhibition or DNA damage, the Cds1dependent checkpoint response may prevent the initiation of new replicons, and/or stabilize existing replicons to prevent replication fork collapse and irreparable damage. It has been pointed out recently that if two replication forks that are converging collapse, this will create a region of unreplicated, and potentially unrepairable, DNA (Elledge 1996). The prevention of such a scenario during perturbations to S phase is certainly an important consideration.

Two models for the DNA-structure checkpoint pathway(s) during S-phase perturbations
In these studies, we have defined a role for Cds1 in an S phase-specific pathway that allows cells to arrest S phase reversibly, and, thus, tolerate the consequence of HU or DNA-damage treatments during S phase. Whereas cds1 null mutants do not show a significant mitotic arrest defect in the first 6 hr when DNA replication is inhibited, the delay seen in these cds1 null cells is unexpectedly dependent on the Chk1 kinase. On the basis of the current data relating to S. pombe DNA structure checkpoints, we propose two models for the action of the checkpoints in response to perturbations in S phase. (Fig.  8A,B).
Model A Cds1 has two separate targets. The first is the replication machinery. Here, Cds1 acts to stabilize Sphase events in response to DNA-replication arrest or DNA damage. The second is the mitotic apparatus. Here, Cds1 targets proteins that regulate p34 cdc2 activity, as has been proposed for Chk1 (Furnari et al. 1997;O'Connell et al. 1997;Peng et al. 1997;Sanchez et al. 1997). If Cds1 is directly transducing the mitotic arrest signal, in its absence the unscheduled activation of Chk1 is obscuring the phenotypic effect.
Model B Cds1 has a single target, the replication machinery, in which Cds1 would act to stabilize S-phase events. To account for the loss of mitotic checkpoint function after HU exposure in cds1.d chk1.d double mutants, we suggest that a, as yet uncharacterized, mechanism links replication directly to p34 cdc2 through a checkpoint Rad-independent pathway. This intrinsic mechanism would, however, require stable replication structures to maintain the link between S phase and mitosis. During normal S phase, this may be active replication itself. During S-phase arrest, the maintenance of S-phase structures would require the checkpoint Rad and Cds1-dependent pathway. An easily conceptualized mechanism for the link between S phase and mitosis in such a model is the sequestration of p34 cdc2 complexes to replication structures, which would keep it unavailable for mitotic initiation. Interestingly, Orp2, Cdc18, and p34 cdc2 may be complexed together during S-phase initiation (Leatherwood et al. 1996;Brown et al. 1997), and Orp2 overexpression is able to arrest the cell cycle independently of the checkpoint rad pathway (Leatherwood et al. 1996).
At present, either variation of our model is consistent with the reported data. One important prediction of the first model is that there is no intrinsic mechanism that ensures the alternation of S phase and M phase in unperturbed cells, because Cds1 (and probably Rad53) activity is not detected during normal S phase. In yeast, there are no published data that allow us to conclude whether such an intrinsic checkpoint is operating. In mammalian cells, however, fusing S phase and G 2 cells shows that normal S phase inhibits mitosis, consistent with an intrinsic checkpoint (Rao and Johnson 1970).
At present, we favor our second model. One reason for this is that it allows for the activity of an intrinsic checkpoint mechanism. Furthermore, it explains an anomaly in the phenotype of the checkpoint rad mutants. When S phase is inhibited in synchronized S-phase cells, mitosis is apparently advanced by ∼30 min relative to untreated cells in rad1, rad3,rad9,rad17,rad26,and hus1 mutants (i.e.,Figs. 1A and 7A). This suggests an intrinsic (checkpoint Rad independent) signal is being lost. This effect is not seen in cdc2.3w cells (Enoch and Nurse 1990;Sheldrick and Carr 1993), which are also defective in the mitotic checkpoint, but in a downstream event indepen- Figure 8. Model for the function of Cds1 in the S-phase arrest response. (A) Checkpoint Rad-dependent activation of Cds1 by perturbations to S-phase acts to ensure both S-phase arrest, by phosphorylating components of the replication complex, and mitotic arrest by phosphorylating a substrate that leads to inactivation of the mitotic apparatus. (B) Cds1 only acts to phosphorylate and stabilize the replication machinery. This prevents inappropriate DNA damage and maintains the structure(s) that are activating a Cds1 independent mitotic arrest signal. This signal would also be independent of the checkpoint Rad proteins, until such time as replication was perturbed, when maintenance of the appropriate replication structures becomes a prerequisite. dent of the effects within S phase. An important consequence of the second model is that it determines that the activity of the checkpoint Rad proteins does not have a direct role in the intrinsic checkpoint mechanism linking S phase and mitosis. The checkpoint Rad proteins would function as ancillary replication proteins required to coordinate S phase only when S phase is perturbed.
At first sight, the checkpoint Rad proteins are directly involved in the DNA-damage checkpoint, which acts through Chk1. Recent evidence in S. cerevisiae, however, clearly shows that the downstream effectors of the checkpoint pathway can operate normally in the complete absence of Mec1 and many other checkpoint proteins when a fragment of an unrelated protein is expressed (Pati et al. 1997). This apparent anomaly of a checkpoint without the key checkpoint proteins is consistent with our model, and could be explained if the role of the checkpoint Rad proteins in the DNA-damage checkpoint were analogous to what we propose in the DNA-replication checkpoint, that is, they act to stabilize specific structures in order that these are available to an as yet uncharacterized mechanism. In S. pombe, this could be (as an example) the direct association of Chk1 with damage/repair structures.

Genetics, molecular biology, and cell biology techniques
Double mutants were created by standard genetic techniques (Gutz et al. 1974). Cell-survival analysis and checkpoint measurements were performed as described previously (Al-Khodairy et al. 1994). Cells were synchronized by elutriation (Aves et al. 1985), or by a 3 hr temperature shift from 27°C to 35°C of either a cdc25.22 mutant strain or a cdc10.V50 mutant strain.

Antibodies for Cds1
GST-Cds1 protein was expressed in S. pombe, purified on glutathione agarose, and used to immunize rabbits. Sera was tested for reactivity against purified protein, S. pombe extracts containing expressed epitope-tagged protein, wild-type S. pombe cell extracts, and cds1 null mutant cells. When compared with preimmune serum, the antibody clearly recognized a Cds1-specific band, which could also be seen in wild-type cells. Several nonspecific bands of approximately the same molecular weight were also seen on Western blots of total wild-type cell extract, even when the antibody was affinity purified against the original antigen. Immunoprecipitation prior to Western analysis abolished these bands, leaving a single Cds1-specific band.

Preparation of protein extract
Logarithmically growing cells were washed in PBS, then washed in lysis buffer [50 mM Tris (pH 7.5), 80 mM ␤-glycerophosphate, 250 mM NaCl, 15 mM nitrophenylphosphate, 50 mM NaF, 5 mM EDTA, 1 mM DTT, and 0.1% NP-40 supplemented with protease inhibitor cocktail (AEBSF, leupeptin, aprotinin, pepstatin, bestatin, and E-64 all at 10 µg/ml final concentration)]. Cells were disrupted with glass beads (BDH), in lysis buffer in a mikrodismembranator (Braun) for 3 × 1 min at 2000 rpm. Protein extract was cleared at 14,000 rpm in a microfuge at 4°C for 5 min and either used fresh or from frozen (LN 2 and storage at −80°C). Preparation of cell extracts from cells expressing tagged proteins from the nmt1 promotor: Cells were grown in the presence of thiamine (promotor repressed), washed several times in distilled water and then inoculated into minimal medium lacking thiamine with the relevant selection. Cells were grown in the absence of thiamine (promoter active) for 18 hr.

Immunoprecipitation of Cds1 and immunoblotting
One milligram of total protein was diluted to 500 µl with lysis buffer and incubated with Cds1 affinity-purified antibody at a dilution of 1:250 at 4°C for 2 hr. Immunocomplexes were collected with protein A agarose (Sigma) for 1 hr at 4°C with mixing. Pellets were washed five times with lysis buffer, boiled in SDS sample buffer, and loaded onto 10% [200:1 acrylamide/bis acrylamide (National Diagnostics)], SDS-polyacrylamide gels. Proteins were transferred to nitrocellulose (MSI). The membranes were blocked with Blotto (PBS, 1% fat free milk powder, 0.05% Tween 20) and incubated in Blotto plus Cds1 antisera (1:1000 dilution), washed in Blotto, and incubated with peroxidase-conjugated secondary antibody (1:5000 dilution. DAKO). The procedure for coimmunoprecipitations was essentially the same as above, except 1 µl of anti-HA (BaBco) was used per sample. Anti c-Myc at a dilution of 1:1000 (Pharmingen) was used to probe the subsequent Western blots. Chemiluminescent detection of HRP-conjugated secondary antibodies was carried out by mixing 10 ml of 100 mM Tris at pH 8.5, 5.4 mM H 2 O 2 with 25 µl of 90 mM p-coumaric acid (Sigma) and 50 µl of 250 mM luminol (Fluka).

Cds1 kinase assay
Alternatively, immunoprecipitates were processed for Cds1 kinase activity. Protein A pellets were washed three times in lysis buffer and three times in kinase buffer (10 mM HEPES at pH 7.5, 75 mM KCl, 5 mM MgCl 2 , 0.5 mM EDTA, 1 mM DTT). Twenty microliters (50% slurry) of bead pellet was incubated with 10 µl of ×2 kinase buffer, 5 µCi of ␣-32 P]ATP (ICN), 1 µl 2 mM ATP, 5 µl of myelin basic protein (1 mg/ml of stock) at 30°C for 15 min. The reaction was stopped by the addition of 20 µl of 2× SDS sample buffer. Samples were run on 15% polyacrylamide gels, fixed in 40% methanol, 10% acetic acid, and dried down, before exposure to film (Hyperfilm, Amersham). Kinase assays were also carried out on GST and 2myc6His-Cds1 fusion proteins. Protein expressed from the nmt1 promoter was purified on glu-tathione-agarose (Sigma) or nickel NTA agarose (Qiagen). DTT and EDTA were omitted from lysis, and wash buffers used for nickel NTA purification.
The kinase activity observed is Cds1 specific, because a Cds1 kinase dead mutant, when expressed in cds1 null cells, does not have appreciable kinase activity when compared with similar experiments with wild-type Cds1. During the production of the cDNA by PCR (primers 5Ј-GAGAGACATATGGAAGAA-CCA-3Ј and 5Ј-AGAGAGGTCGACACTCGA-3Ј) and the site directed mutagenesis to produce the kinase dead allele cds1.KD (Primer 5Ј-GCCAAGCCAAATTCAGATATTTTAAGATGG-3Ј), we sequenced the constructs to avoid unwanted mutations. During this analysis, we identified two separate changes compared with the published sequence. These were verified as correct by sequencing the same region of the genomic clone. The modified sequence, including introns, of the cds1 gene has been submitted to the database with the accession no. AJ222869.

Phosphatase treatment
Immunoprecipitates were washed three times in lysis buffer and three times in calf alkaline phosphatase (CIAP) buffer (50 mM Tris at pH 8.0, 5 mM MgCl 2 ), treated with the indicated units of CIAP (GIBCO BRL) for 30 min at 37°C and then processed for Western blotting.

S-phase checkpoint analysis
Method 1 Early G 2 cells from log-phase cultures of cyr1 sxa2 strains were collected by centrifugal elutriation. After 30 min incubation in minimal media at 30°C, P-factor was added to a final concentration of 1µg/ml. Samples were scored for septation every 30 min. After the peak of septation, the culture was divided in two. One-half was treated with 0.033% MMS (final concentration), and cultures were incubated for 30 min at 30°C. Cells were harvested by centrifugation, washed three times with water to release them from the P-factor-induced G 1 arrest, resuspended in minimal media or minimal media + 0.033% MMS, and incubated for 4 hr at 30°C. Samples were taken at the time of release and then every 30 min and fixed in 70% EtOH (Sazer and Sherwood 1990). Fixed cells were treated with RNase, stained with propidium iodide, and analyzed for DNA content with a Coulter Epics Elite flow cytometer.
Method 2 For starvation-induced synchronization in G 1 , logphase cells grown in minimal media were harvested, washed three times with water, resuspended in minimal media lacking a nitrogen source and incubated 18 hr at 27°C. Afterward, cells were harvested, washed with water, and allowed to recover by incubation in complete media at 30°C for 1.5 hr. The cells were then harvested, washed twice in water, and divided into two cultures in minimal media, one containing 0.033% MMS. The cultures were incubated at 30°C and samples were taken every half hour and fixed in 70% EtOH. Fixed cells were processed as described above including one step of sonication and analyzed for DNA content with a Becton-Dickinson FACScan flow cytometer.


extfieldsofstudy: Chemistry Medicine

text:
Tyrosine Structural Changes Detected during the Photoactivation of Rhodopsin*

We present the first Fourier transform infrared (FTIR) analysis of an isotope-labeled eukaryotic membrane protein. A combination of isotope labeling and FTIR difference spectroscopy was used to investigate the possible involvement of tyrosines in the photoactivation of rhodopsin (Rho). Rho → MII difference spectra were obtained at 10 °C for unlabeled recombinant Rho and isotope-labeledl-[ring-2H4]Tyr-Rho expressed in Spodoptera frugiperda cells grown on a stringent culture medium containing enrichedl-[ring-2H4]Tyr and isolated using a His6 tag. A comparison of these difference spectra revealed reproducible changes in bands that correspond to tyrosine and tyrosinate vibrational modes. A similar pattern of tyrosine/tyrosinate bands has also been observed in the bR → M transition in bacteriorhodopsin, although the sign of the bands is reversed. In bacteriorhodopsin, these bands were assigned to Tyr-185, which along with Pro-186 in the F-helix, may form a hinge that facilitates α-helix movement.

Thus far, bR is the only IMP with a seven-helix transmembrane motif whose structure has been elucidated at atomic resolution (17)(18)(19). This structure confirmed key features of an earlier "spectroscopic" model based in part on site-directed mutagenesis and FTIR difference spectroscopy (20 -23). For example, a retinal binding pocket was predicted on the basis of FTIR, UV-visible spectroscopy, and site-directed mutagenesis (20 -24), which is formed in part from several residues on the F-helix (helix 6 in rhodopsin), including two tryptophans (Trp-182, Trp-189), a proline (Pro-186), and a tyrosine (Tyr-185). These residues, along with several others from the C-helix (Trp-86, Thr-89, and Asp-85) are in close proximity to the retinal chromophore and act to constrain its structure in an all-trans configuration. In addition, these residues are in a good position to couple retinal isomerization to protein changes involved in proton transport, including a change in the structure and orientation of the F-helix.
A comparable combination of Trp, Pro, and Tyr residues (WXPY) is fully conserved in helix 6 of all visual pigments except the blue pigments where it is YXPY. It is possible that, although the tyrosine-proline sequence is reversed, a similar coupling between chromophore isomerization and protein exists in the two systems (22) despite their different functions and evolutionary pathways. To investigate this possibility, we have used for the first time a combination of FTIR difference spectroscopy and isotope labeling to analyze tyrosine structural changes in rhodopsin. A comparison with earlier results obtained for bacteriorhodopsin reveals that a similar, although reversed, pattern of tyrosine/tyrosinate changes does occur. This reversed pattern might be explained if the all-trans to cis transition of the chromophore in both systems is coupled to similar changes in adjacent residues in agreement with an earlier suggestion (25).

EXPERIMENTAL PROCEDURES
Preparation of Isotope-labeled Rhodopsin-Wild-type hexahistidinetagged rhodopsin derived from bovine rhodopsin was produced and purified as described previously (26). The hexahistidine tag introduces a slight redshift in the max of rhodopsin from 498 to 500 nm. For the purpose of isotope labeling, Spodoptera frugiperda (Sf9) cells (ATCC: CRL-1711) were grown and subsequently infected in a customized serum-free and protein-free medium partially depleted in L-Tyr (Biowhittaker, Walkersville, MD) to which an additional 2 mM L-[ring-2 H 4 ]Tyr (Cambridge Isotope Laboratories, Andover, MA) was added. To determine label incorporation levels, total proteoliposome fractions were treated with Pronase to generate free amino acids. Subsequent gas chromatography-mass spectrometry analyses of two different samples revealed that 70 Ϯ 3% of membrane protein-incorporated tyrosines were labeled. 2 No detectable labeling was found to occur in Phe residues as expected, because in eukaryotic cells Tyr is degraded through the Krebs cycle, and thus the labels should not be incorporated into an essential amino acid like Phe.
FTIR Spectroscopy-Rhodopsin films for transmission IR spectroscopy were prepared by isopotential spin-drying (5, 27) of a 1-ml aqueous suspension of membranes containing approximately 3 nmol of unlabeled or L-[ring-2 H 4 ]Tyr-labeled Rho-His 6 onto a AgCl window. The film is then rehydrated prior to insertion into a sealed transmittance cell as described previously (28). Transmission FTIR difference spectra of the hydrated rhodopsin films were recorded at 10°C using methods similar to those reported previously (29,30). Briefly, the H 2 O content of the sample was monitored by measuring the intensity ratio of the 3400cm Ϫ1 band (O-H stretch mode) to the protein Amide II band near 1545 cm Ϫ1 . The Rho 3 Meta II difference spectra were obtained as follows: the sample was photobleached for 3 min using light from a 150-watt tungsten illuminator (model 180, Dolan-Jenner Industries, Lawrence, MA) filtered by a 500-nm long-pass filter (Corion Corp., Holliston, MA) and several heat filters and transmitted to the sample with an annular optical fiber. Spectra were recorded at 8 cm Ϫ1 resolution and 5-min intervals for several hours before and after illumination (1350 scans for each spectrum) on a Bio-Rad FTS-60A spectrometer (Bio-Rad, Digilab Division, Cambridge, MA) equipped with a mercury-cadmium-teluride detector. Each difference spectrum shown represents a subtraction of the spectrum recorded immediately before the light is turned on from the spectrum recorded immediately after illumination. The larger set of spectra are recorded prior to and after illumination to monitor sample stability, hydration, and Meta II decay.
FTIR transmission spectra of tyrosine and L-[ring-2 H 4 ]Tyr-saturated solutions were measured at room temperature at pH 12. Each sample was prepared by dissolving Tyr or L-[ring-2 H 4 ]Tyr to saturation in a borate buffer that was titrated using KOH to pH 12. A small drop (ϳ7 l) of this solution was placed on a CaF 2 window, and a second CaF 2 window was pressed against it to form a uniform thin film. The windows were inserted into a liquid sample cell (Harrick Scientific Corp., Ossining, NY; TFC-M32) and mounted in a Nicolet 740 FTIR spectrometer (Nicolet Instruments, Madison, WI) equipped with a triglyceride sulfate detector. Spectra were collected consisting of 512 scans. An identical method was used to record a spectrum of the buffer, which was interactively subtracted from the absorption spectrum of the sample using the 3400-cm Ϫ1 water band as a reference. Fig. 1 shows a comparison of the Rho 3 Meta II difference spectra from WT and L-[ring-2 H 4 ]Tyr-Rho in the region from 1800 to 1000 cm Ϫ1 . To assess the reliability of assignments using tyrosine isotope labeling, three samples of both WT and L-[ring-2 H 4 ]Tyr-Rho were measured independently, and the resulting data are shown in Fig. 1. All spectra show very similar features, reflecting changes of the retinylidene chromophore, which undergoes an 11-cis to all-trans conformational transition, as well as structural changes of the protein required to create a binding site for the G-protein transducin. For example, negative bands have been assigned to the Rho chromophore on the basis of comparison with results from resonance Raman spectroscopy at 1548 cm Ϫ1 (Ϫ) (CϭC ethylenic stretch) and 1238 cm Ϫ1 (Ϫ) (C 12 -C 13 stretch mode) 3 (31)(32)(33)(34). In contrast, few bands have been assigned thus far to specific amino acids except in the 1700 -1800-cm Ϫ1 region (CϭO stretch modes of Asp and Glu carboxylic acid groups (28,35)) and near 2550 cm Ϫ1 (the SH stretch mode of cysteine residues (36)).

RESULTS
As seen in Fig. 1, the introduction of L-[ring-2 H 4 ]Tyr into Rho produces several highly reproducible changes in the region between 1800 and 1000 cm Ϫ1 . In particular, several bands are altered with a change in intensity, reflecting an isotope-induced shift in frequency, including those near 1595, 1570, 1517, 1423, 1373, 1348, 1275, 1253, 1221 (sh), and 1178 cm Ϫ1 . There are also bands near 1643 and 1535 cm Ϫ1 whose intensities are not reproducible within each set and are therefore not assigned to tyrosine vibrations. This variability is most likely due to sample instability observed in the highly absorbing amide I and II regions as well as small differences in sample preparation which can give rise to polarization effects.
In general, the changes we have assigned to tyrosine vibrations can be caused either by an isotope-induced frequency shift of a tyrosine/tyrosinate vibrational mode out of a particular region or by a shift of a labeled tyrosine/tyrosinate mode into that region of the spectrum. Detailed assignments can be made on the basis of comparison with Tyr and L-[ring-2 H 4 ]Tyr model compounds. For example, a decrease in intensity at 1595 cm Ϫ1 and increase at 1570 cm Ϫ1 due to L-[ring-2 H 4 ]Tyr incorporation (Fig. 1, inset) agrees well with the isotope induced downshift of the 1598 cm Ϫ1 ring stretching mode of tyrosinate to 1569 cm Ϫ1 in L-[ring-2 H 4 ]Tyr measured at pH 12 (Fig. 2). Similarly, a drop in intensity near 1275 cm Ϫ1 and increase near 1253 cm Ϫ1 (Fig.  1, inset) (this increase appears as a maximum at 1245 cm Ϫ1 in a double difference spectrum; not shown) agrees qualitatively with the isotope-induced shift of a band at 1271 cm Ϫ1 in Ltyrosinate to 1238 cm Ϫ1 in L-[ring-2 H 4 ]Tyr (Fig. 2). Previously this band was assigned to the C-O Ϫ stretch mode of tyrosinate (37,38). Overall, these data, and in particular the appearance of positive bands characteristic of (labeled) tyrosinate, indicate that a tyrosine residue is partially deprotonated in the Meta II state. This might reflect a partial deprotonation of a tyrosine residue in Rho upon formation of Meta II (see below) or, alternatively, a change in environment of an existing tyrosinate in Rho upon Meta II formation.
Additional bands that exhibit reproducible isotope-induced changes are assignable to vibrational modes found in tyrosine or L-[ring-2 H 4 ]Tyr. However, the pattern is complex and may reflect one or more tyrosines that undergo a change in environment or protonation state during Meta II formation. For example, the positive 1517-cm Ϫ1 band, which drops in intensity, is close to the frequency of an intense tyrosine ring vibration in L-Tyr (37,39) and therefore reflects a tyrosine in the Meta II state. Other bands that match known tyrosine frequencies are at 1373 cm Ϫ1 (not assigned), 1250 cm Ϫ1 (bending mode of tyrosine hydroxyl group), and 1179 cm Ϫ1 (ring CH bending mode). The 1373-and 1179-cm Ϫ1 bands are positive and therefore most likely reflect tyrosine(s) in Meta II. The 1373-cm Ϫ1 band shifts to 1348 cm Ϫ1 in L-[ring-2 H 4 ]Tyr-Rho in agreement with published spectra (38). However, the increased intensity at 1250 cm Ϫ1 may reflect a change in a tyrosine residue in the unbleached state of Rho. Part of the intensity changes in this region were already attributed to the downshift of a positive tyrosinate band from 1277 cm Ϫ1 to near 1248 cm Ϫ1 in the Meta II state (see above). However, a prominent tyrosine mode due to the bending vibration of the tyrosine hydroxyl also occurs at this frequency and downshifts in L-[ring-2 H 4 ]Tyr to 1226 cm Ϫ1 (38) in good agreement with the change observed at 1221 cm Ϫ1 in L-[ring-2 H 4 ]Tyr-Rho. This band is not likely to be due to residual Meta I, which exhibits a band near 1215 cm Ϫ1 (30, 40), because no increase in the corresponding Meta I specific band at 950 cm Ϫ1 is observed (not shown).
Finally, a prominent drop in intensity at 1423 cm Ϫ1 (Fig. 1) in L-[ring-2 H 4 ]Tyr-Rho might be due to the isotope-induced downshift of negative tyrosine ring mode from the 1515 cm Ϫ1 region. A band due to such a ring mode has been previously identified in the L-[ring-2 H 4 ]Tyr-labeled Photosystem II in the Tyr D to Tyr D* FTIR difference spectrum (38). Alternatively, this change might be due to a mode observed at 1418 cm Ϫ1 in L-[ring-2 H 4 ]Tyr at high pH, which downshifts from 1498 cm Ϫ1 in L-Tyr at high pH (Fig. 2). This would indicate that a (partially) ionized tyrosine exists in Rho. However, the loss of the corresponding unlabeled tyrosinate vibration at 1498 cm Ϫ1 would lead to an increase in intensity near 1498 cm Ϫ1 in L-[ring-2 H 4 ]Tyr-Rho, which is not clearly observed, but could be partially masked by the loss of the positive band at 1517 cm Ϫ1 .
Strikingly, there is a similar pattern of tyrosinate-related spectral changes observed in the bR 3 M difference spectrum of bR (41) and the Rho 3 Meta II difference spectrum of Rho when L-[ring-2 H 4 ]Tyr is incorporated, although the sign of the bands is reversed. Similar to the results reported above for Rho, L-[ring-2 H 4 ]Tyr labeling causes a downshift of a band from 1590 to 1570 cm Ϫ1 in the bR difference spectrum (Fig. 3, inset), although the downshifted bands are positive in the Rho spectrum and negative in the bR spectrum. Also, similar to rhodopsin a band at 1276 cm Ϫ1 appears to shift to near 1248 cm Ϫ1 in the bR 3 M difference spectrum (Fig. 3, inset). Additional peaks were assigned to tyrosine vibrations previously in the bR 3 M difference spectrum of bR (41) at 1517 cm Ϫ1 (Ϫ) (tyrosine) and 1419 -1410 cm Ϫ1 (ϩ) (L-[ring-2 H 4 ]tyrosine). Here again, the frequencies are similar to bands observed in the Rho 3 Meta II difference spectrum, although the signs of the bands are reversed. DISCUSSION FTIR difference spectroscopy has been used extensively to study conformational changes in membrane proteins (42)(43)(44). In the case of rhodopsin, structural changes of the retinylidene chromophore and protonation and/or hydrogen bonding changes of Asp, Glu, and Cys residues have been detected at  different stages of the photoactivation cascade (28,30,35,36,(45)(46)(47)(48)(49)(50)(51). In some cases, assignment of vibrational bands to individual amino acid residues in rhodopsin was facilitated by site-directed mutagenesis (28,35). FTIR difference spectroscopy can also be used to study changes in the environment of individual water molecules in rhodopsin during the different steps in the photoactivation cascade (52)(53)(54).
In this work, FTIR difference spectroscopy and amino acid isotope labeling were combined for the first time to analyze structural changes of tyrosines in rhodopsin. Isotope labeling was accomplished by growing Sf9 insect cells on a stringent medium containing L-[ring-2 H 4 ]Tyr. Tyrosine bands can be identified and assigned on the basis of the isotope induced shifts and by comparison with a series of model tyrosine compounds at high and low pH. Our results clearly demonstrate that a set of bands in the Rho 3 Meta II difference spectra can be assigned to vibrational modes of tyrosine and tyrosinate. On this basis, it can be concluded that one or more tyrosines participate in the rhodopsin photocascade up to formation of the Meta II intermediate and that at least a partial protonation of a tyrosinate is involved. A possible involvement of tyrosine residues was previously proposed on the basis of UV difference spectroscopy (55) and site-specific mutagenesis (56) and also suggested by a recent study using UV resonance Raman spectroscopy (57).
The similarity between the pattern of isotope-induced changes observed in bacteriorhodopsin and rhodopsin is particularly interesting. Earlier studies based on tyrosine isotope labels (41), site-directed mutagenesis (21), and site-directed isotope labeling (58) concluded that tyrosine bands identified in the FTIR difference spectra of bacteriorhodopsin arose from protonation changes in Tyr-185 located in the F-helix. Subsequent FTIR measurements on bR containing a 13 C isotope label in the C1 position of Tyr-185 indicated that the Tyr-185/Pro-186 peptide bond buried in the center of the F-helix undergoes some type of structural rearrangement (59). An interesting possibility is that the Tyr-185/Pro-186 bond serves as a hinge, which gives rise to the apparent tilting of the F-helix late in the bR photocycle (60,61).
The present findings indicate that a similar, although reversed, pattern of tyrosine/tyrosinate changes occurs also upon photoactivation of rhodopsin. Although on the basis of this study we cannot assign these changes to any particular tyrosine(s) in rhodopsin, an interesting candidate is Tyr-268. This residue is also positioned adjacent to a Pro residue (Pro-267). It is well known that Pro residues break ␣-helical structure and can function as kinks allowing independent movement of adjacent helical segments. The existence of a Tyr/Pro pair in the center of helix 6 in bacteriorhodopsin (Tyr-185, Pro-186) and a Pro/Tyr pair in rhodopsin (Pro-267, Tyr-268) raises the possibility that such a pair acts synergistically independent of the order of these residues, thereby forming a potential hinge for movements in helix 6 in response to chromophore isomerization. Indeed, several lines of evidence have been presented that a rigid body movement involving part of helix 6 takes place during photoactivation of rhodopsin (62)(63)(64). Also, recent structural models for rhodopsin (e.g. Refs. 64 and 65) place Tyr-268 close to Glu-113, suggesting a possible role in the deprotonation of the Schiff base upon Meta II formation, which could be related to the (partial) protonation indicated by our data. In fact, mutagenesis studies show that replacement of Tyr-268 by Phe perturbs ligand binding and activation of rhodopsin (56) and that the adjacent Trp-265 also is involved structurally in formation of Meta II (66, 67). Actually, this region probably is one of the hot spots in rhodopsin activation, because in the vicinity of Tyr-268 are also residues Gly-121 and Phe-261, which form a synergistic functional unit with the 9-methyl group of the chromophore (68). Mutation of these residues in most cases has a profound effect on dark and/or rest activity of rhodopsin (68,69). However, to definitely identify Tyr-268 as (one of) the residue(s) responsible for the changes we observe additional site-specific mutagenesis and/or site-directed isotope labeling will be required. Because mutagenesis tends to perturb functionally or structurally active sites, site-directed isotope labeling will be the preferred method.
Our results suggest a common element for the primary mechanism of rhodopsin and bacteriorhodopsin that goes beyond the general features of a retinylidene chromophore and a seven-helix transmembrane domain. In particular, it establishes that similar tyrosine/tyrosinate changes occur during activation of both proteins. In fact, the reversal of the signs of tyrosine/tyrosinate assigned bands may be related to the respective role of the all-trans chromophore in the activation of the rhodopsin and bacteriorhodopsin systems as indicated by FTIR studies on the effects of removal of the all-trans chromophore in rhodopsin (29) and bacteriorhodopsin (25). If these similarities arise in changes in the F-helix (helix 6), then it is possible that a common mechanism of chromophore-protein coupling exists in both proteins. This is a very important issue that will be addressed by subsequent studies using a combination of FTIR difference spectroscopy, mutagenesis, and isotope labeling.