The Plant Cell, Vol. 16, 857–873, April 2004, www.plantcell.org ª 2004 American Society of Plant Biologists
Purification and Characterization of Serine Proteases That
Exhibit Caspase-Like Activity and Are Associated with
Programmed Cell Death in Avena sativa
Warren C. Coffeen and Thomas J. Wolpert1
Department of Botany and Plant Pathology and Center for Gene Research and Biotechnology, Oregon State University,
Corvallis, Oregon, 97331-2902
Victoria blight of Avena sativa (oat) is caused by the fungus Cochliobolus victoriae, which is pathogenic because of the
production of the toxin victorin. The victorin-induced response in sensitive A. sativa has been characterized as a form of
programmed cell death (PCD) and displays morphological and biochemical features similar to apoptosis, including
chromatin condensation, DNA laddering, cell shrinkage, altered mitochondrial function, and ordered, substrate-specific
proteolytic events. Victorin-induced proteolysis of ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) is shown to
be prevented by caspase-specific and general protease inhibitors. Evidence is presented for a signaling cascade leading to
Rubisco proteolysis that involves multiple proteases. Furthermore, two proteases that are apparently involved in the
Rubisco proteolytic cascade were purified and characterized. These proteases exhibit caspase specificity and display
amino acid sequences homologous to plant subtilisin-like Ser proteases. The proteases are constitutively present in an
active form and are relocalized to the extracellular fluid after induction of PCD by either victorin or heat shock. The role of
the enzymes as processive proteases involved in a signal cascade during the PCD response is discussed.
INTRODUCTION
Victoria blight of Avena sativa (oat) is caused by the necrotrophic
fungus, Cochliobolus victoriae (Meehan and Murphy, 1946),
which is pathogenic because of the production of the hostspecific toxin, victorin (Meehan and Murphy, 1947). Isolates of
C. victoriae that produce victorin are pathogenic on susceptible
A. sativa (Meehan and Murphy, 1947), whereas mutants or outcrosses that do not produce the toxin are nonpathogenic. Host
susceptibility and victorin sensitivity are conferred by a dominant
allele at the Vb locus (Litzenberger, 1949). Homozygous recessive genotypes (vb/vb) are victorin insensitive and resistant
to the fungus. The interaction between victorin and the Vb gene
product induces a response in A. sativa that displays characteristics of programmed cell death (PCD) (Navarre and Wolpert,
1999; Tada et al., 2001; Yao et al., 2001; Curtis and Wolpert,
2002, 2004).
PCD is a genetically controlled, organized form of cellular
suicide that functions in eliminating unnecessary or aged cells. It
is essential for cellular maturation and morphogenesis and is
required to maintain cellular homeostasis in multicellular organisms. In addition, improper regulation of PCD has been impli-
1 To
whom correspondence should be addressed. E-mail wolpertt@
bcc.orst.edu; fax 541-737-3573.
The author responsible for distribution of materials integral to the
findings presented in this article in accordance with the policy described
in the Instructions for Authors (www.plantcell.org) is: Thomas J. Wolpert
([email protected]).
Article, publication date, and citation information can be found at
www.plantcell.org/cgi/doi/10.1105/tpc.017947.
cated in a wide variety of animal diseases (Polverini and Nör,
1999; Wang and Wang, 1999).
PCD also has been associated with several processes in
plants, including senescence (Bleecker and Patterson, 1997;
Miller et al., 1999; Schmid et al., 2001), stress (Katsuhara, 1997;
Solomon et al., 1999), development (Runeberg-Roos and
Saarma, 1998; Groover and Jones, 1999; Schmid et al., 1999),
and the hypersensitive response (HR) to pathogens (Dangl et al.,
1996; Mittler et al., 1997; Pontier et al., 1998; Mackey et al., 2002;
Abramovitch et al., 2003). Currently, very little is known about the
fundamental processes that control and regulate PCD in plants.
Apoptosis, the most characterized form of PCD, has been
extensively studied in animal systems and can be distinguished
by unique characteristics. Cells undergoing apoptosis display
morphological changes, including cell shrinkage, chromatin
condensation, and apoptotic body formation. Biochemically,
apoptotic cells exhibit DNA fragmentation (also referred to as
DNA laddering) and activation of a family of Cys proteases
called caspases (cysteine aspartases) (reviewed in Vaux and
Korsmeyer, 1999; Hengartner, 2000). The name caspase is
derived from their active site residue (which is a Cys) and substrate specificity (caspases cleave only after aspartate residues, aspase).
The victorin-induced PCD response in sensitive A. sativa
demonstrates similar morphological and biochemical traits to
animal apoptosis, including cell shrinkage and collapse (Yao
et al., 2001; Curtis and Wolpert, 2004), chromatin condensation
(Yao et al., 2001), DNA laddering (Navarre and Wolpert, 1999;
Tada et al., 2001), mitochondrial depolarization and permeability
transition (Curtis and Wolpert, 2002, 2004), and ordered, substrate-specific proteolytic events (Navarre and Wolpert, 1999).
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Furthermore, victorin-induced PCD in A. sativa is easily initiated,
proceeds in a rapid and synchronous manner, and appears to
encompass at least all leaf mesophyll cells. Therefore, victorin
treatment of A. sativa provides an appropriate system in which to
study the mechanism and progression of plant PCD.
The proteolytic cleavage of the large subunit of ribulose-1,5bisphosphate carboxylase/oxygenase (Rubisco) has been identified as a specific PCD-induced event in victorin-treated
A. sativa cells (Navarre and Wolpert, 1999). Rubisco cleavage
occurs after the first 14 amino acids (apparently after glutamate14) and is prevented by Cys protease inhibitors (Navarre and
Wolpert, 1999). Rubisco degradation has been identified as
a characteristic of senescence (Weidhase et al., 1987; Ferreira
and Davies, 1989), also a form of PCD, and has been shown to
occur in A. sativa chloroplast after oxidative stress (Casano and
Trippi, 1992), a treatment shown to induce PCD (Amor et al.,
1998; Solomon et al., 1999). Additionally, chloroplast-localized
proteases have been reported that appear to recognize Rubisco
as a substrate (Bushnell et al., 1993; Casano et al., 1994). These
data indicate that a specific proteolytic process is required to
degrade Rubisco and that this process is common to several
types of PCD.
Proteolytic alteration of key cellular proteins is a fundamental
characteristic of animal apoptosis and is executed by the highly
specialized caspases. Multiple cellular targets exist for caspases, all of which are directly or indirectly involved in the
ordered disassembly of the cell. Two types of caspases exist,
initiator and effector caspases. Initiator caspases are activated
by autoproteolysis, after which they are able to proteolytically
activate effector caspases. Effector caspases target cellular
proteins, such as poly(ADP-ribose) polymerase (PARP), which
is proteolytically inactivated, resulting in a signature 89-kD
fragment (reviewed in Solary et al., 1998). In addition, CAD, a
caspase-activated DNase, is indirectly activated when an
effector caspase cleaves ICAD (inhibitor of CAD), allowing CAD
to disassociate and process DNA into nucleosomal fragments
(reviewed in Nagata, 2000). Caspase proteolysis typifies the
ordered processive nature of apoptosis because caspases
function with exquisite control, cleaving at limited site(s) found
within specifically targeted key proteins, differing from the
degradative role of other proteolytic pathways required during
apoptosis (e.g., the proteasome-ubiquitin pathway of protein
degradation) (Solary et al., 1998).
Current research has demonstrated that caspase-specific
inhibitors can prevent different forms of plant PCD, suggesting
that caspase-like enzymes may be involved in these responses
(reviewed in Woltering et al., 2002). For example, Tobacco
mosaic virus–induced HR in Nicotiana tabacum leaves was prevented by treatment with short peptide inhibitors that preferentially inhibit caspase-1 and caspase-3 proteases (del Pozo
and Lam, 1998). Other forms of PCD in N. tabacum, including those induced by bacteria (Richael et al., 2001) and
treatment with a fungal elicitor (Elbaz et al., 2002), also were
prevented by caspase inhibitors. Similar results were obtained
with caspase inhibitors in preventing or reducing cell death after
treatment with camptothecin (De Jong et al., 2000), staurosporine (Elbaz et al., 2002), and isopentenyladenosine (Mlejnek and
Procházka, 2002). In addition, heat shock–induced PCD in
N. tabacum suspension cells stimulated cleavage of PARP and
activation of a caspase-3–like protease (Tian et al., 2000),
whereas N. tabacum protoplast treated with menadione displayed PARP cleavage (Sun et al., 1999). However, to date, no
protease that recognizes caspase substrates or is inhibited by
caspase inhibitors has been identified in plants.
Studies have described proteases associated with several types of plant PCD, including senescence (Delorme et al.,
2000; Schmid et al., 2001; Eason et al., 2002), oxidative stress
(Solomon et al., 1999), seed development (Schmid et al., 1998,
1999; Wan et al., 2002), tracheary element development
(Runeberg-Roos and Saarma, 1998; Groover and Jones, 1999),
and the HR (Vera and Conejero, 1988; D’Silva et al., 1998; Krüger
et al., 2002). However, characterization of these proteases
reveals that they are degradative, not processive enzymes; thus,
they are unlike caspases or other proteases involved in signaling
pathways. A degradative nature suggests that their role in PCD
may be associated with the terminal decomposition of the dying
cell but not in the initiation or progression of the PCD signal.
This research describes the purification and characterization
of two proteases that possess caspase-like specificity but contain an active-site Ser residue. Therefore, because of their
aspartate specificity (aspase) and active-site Ser residue, they
have been termed saspases. The saspases, purified from A.
sativa, contain amino acid sequence similarities to subtilisin-like
Ser proteases from plants and likely function in a PCD-induced
signaling cascade involving other proteases that leads to the
proteolytic processing of Rubisco. They also are constitutively
present in the cell, not being transcriptionally or translationally
activated during the response, but released into the extracellular
fluid (ECF) upon induction of PCD. Heat shock–induced PCD
also is further characterized and displays similar biochemical
features as victorin-induced PCD, including DNA laddering,
Rubisco proteolysis, and release of the saspases into the ECF.
These results and the implications of numerous subtilisin-like Ser
proteases involved in plant disease, development, and death are
discussed.
RESULTS
Evidence for a Protease Cascade Leading to
Rubisco Proteolysis
Rubisco proteolysis is a characteristic of victorin-induced PCD
and has been shown to be inhibited by two general protease
inhibitors, E-64 (a Cys protease inhibitor) and leupeptin (a Cys
and Ser protease inhibitor) (Navarre and Wolpert, 1999). Two
caspase and a granzyme B inhibitor were tested for their effects
on Rubisco proteolysis. These and a large variety of other short
peptide inhibitors (and corresponding substrates) were designed
based on the caspase recognition sites in known substrates.
These inhibitors are typically preferential for a given caspase but
can often inhibit a wide variety of caspases (Kidd, 1998). One
inhibitor initially evaluated, Ac-VAD-CMK, is considered a
pan-caspase inhibitor because it can inhibit most caspases
(Thornberry et al., 1997; Kidd, 1998; Ekert et al., 1999), whereas
another inhibitor, Ac-DEVD-CMK, is more specific for caspase-3 (Thornberry et al., 1997; Ekert et al., 1999). In addition,
Caspase-Like Ser Proteases
we tested Z-AAD-CMK, which inhibits granzyme B (Ekert et al.,
1999), a Ser protease that specifically cleaves peptide bonds
after aspartate residues (the recognition sequence for the three
inhibitors is in single-letter amino acid code). Preincubation of
peeled A. sativa leaves with the caspase and granzyme B inhibitors prevented victorin-induced Rubisco proteolysis (Figure 1).
The effectiveness of these inhibitors was similar to that of E-64
and leupeptin.
The inhibition of Rubisco proteolysis by caspase-specific
inhibitors suggested that a caspase-like protease(s) might be
present in victorin-treated A. sativa tissue. Total soluble protein
was extracted from 60 g of victorin-treated A. sativa leaves, and a
low level of proteolytic activity that hydrolyzed the pan-caspase
substrate Z-VAD-AFC was observed (data not shown). To concentrate the activity, the soluble protein extract was run through
hydrophobic interaction chromatography (HIC) (see Methods).
Two distinct proteolytic activities were separated during chromatography: activity A (Figure 2A, fractions 56 to 67), which
hydrolyzed Z-DEVD-AFC (a caspase-3 substrate) and eluted at
0.45 M ammonium sulfate, and activity B (Figure 2A, fractions
76 to 100), which hydrolyzed Z-VAD-AFC and eluted at 0.3 M
ammonium sulfate. The fractions containing each activity were
individually combined for further analysis. Activity A hydrolyzed
the substrate Z-DEVD-AFC but not Z-VAD-AFC or Z-AAD-AFC (a
substrate for granzyme B) (Figure 2B), whereas activity B
hydrolyzed Z-VAD-AFC and Z-AAD-AFC but not Z-DEVD-AFC
(Figure 2C). Figure 2D illustrates the percentage of inhibition of
activities A and B by the protease inhibitors used to prevented
Rubisco proteolysis in Figure 1. Z-VAD-AFC hydrolysis by
activity B was completely inhibited by the caspase inhibitors
Ac-VAD-CMK and Z-AAD-CMK but was unaffected by AcDEVD-CMK, leupeptin, and E-64 (Figure 2D, open bars). However, activity A hydrolysis of Z-DEVD-AFC was only inhibited by
Ac-DEVD-CMK (Figure 2D, closed bars). These data suggest
that there are two distinct caspase-like protease activities
involved in the proteolytic processing of Rubisco because
granzyme B and both caspase-specific inhibitors all prevent
Rubisco proteolysis in vivo, whereas two of the inhibitors inhibit
only activity B, and the other inhibits only activity A. Furthermore,
because neither caspase-like activity is inhibited by leupeptin or
E-64, both of which prevent Rubisco proteolysis, a third noncaspase-like protease is also likely involved. Thus, a proteolytic
cascade involving at least three proteases, two caspase-related
and a non-caspase-like protease, is apparently activated during
Figure 1. Inhibition of Rubisco Proteolysis with Protease Inhibitors.
Leaf segments with epidermis removed were pretreated with 200 mM of
the indicated inhibitor for 2 h then with 20 ng/mL of victorin for 4 h.
Coomassie blue–stained 12% SDS-PAGE of total protein extract. Only
the region containing Rubisco is shown.
859
Figure 2. Characterization of Two Caspase-Like Proteolytic Activities.
(A) Activity profile of fractions from a HIC column. The protease in activity
A was assayed by incubating 50 mL of each fraction with 50 mL of 20 mM
Mops, pH 7, with 20 mM substrate for 2 h. Activity B was assayed as
described in Methods. RFU, relative fluorescence units.
(B) Substrate profile of activity A. Activity A was assayed as described
above for proteolytic activity with indicated substrates. RFU, relative
fluorescence units.
(C) Substrate profile of activity B. RFU, relative fluorescence units.
(D) Inhibitor profile of activities A and B. Samples were pretreated with
inhibitor (200 mM) for 2 h and then assayed for hydrolytic activity with
Z-DEVD-AFC (activity A) or with Z-VAD-AFC (activity B). Expressed
as percent inhibited of noninhibited control.
victorin-induced PCD, leading to the proteolytic processing of
Rubisco.
Reversible and Irreversible Inhibition by
Caspase Inhibitors
The protease(s) constituting activity B from the HIC column,
which hydrolyzed Z-VAD-AFC and Z-AAD-AFC, displayed the
highest levels of activity; therefore, this activity was selected for
further characterization. No further characterization of activity A,
which hydrolyzed Z-DEVD-AFC, was performed in this study.
Initial attempts to visualize the activity B protease with the
biotinylated caspase inhibitor biotin-VAD-FMK on a biotin blot
were unsuccessful. Biotin-VAD-FMK prevented Z-VAD-AFC hydrolysis but did not covalently label the protein (data not shown).
Three types of inhibitors with the same tripeptide recognition
motif, Ac-VAD-CMK, Z-VAD-FMK, and Ac-VAD-CHO, were
tested for the nature of their binding to the protease (Figure
3A). Chloromethylketone (CMK)-based inhibitors irreversibly
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The biotinylated inhibitor biotin-YVAD-CMK irreversibly binds
and inhibits the activity of the saspase (activity B). However,
because of the highly reactive nature of CMKs (Ekert et al., 1999),
biotin-YVAD-CMK labeled numerous proteins in the fractions
containing activity B from the HIC column. To determine if
labeling of any of these proteins was specific, that is, if biotinYVAD-CMK binds to the active site of one of these proteins, a
nonbiotinylated inhibitor that also inhibits saspase activity (binds
to the active site) was preincubated with the protein sample to
identify proteins whose biotin-YVAD-CMK binding was outcompeted/blocked. Preaddition of Ac-VAD-CMK outcompeted
binding of biotin-YVAD-CMK to an 84-kD protein (Figure 3B, lane
2) but did not affect labeling of any other proteins (data not
shown). Conversely, preaddition of Ac-DEVD-CMK, which does
not inhibit saspase (activity B) activity, did not block binding of
biotin-YVAD-CMK to the 84-kD protein (Figure 3B, lane 3).
Furthermore, other inhibitors that inhibit saspase activity (e.g.,
Z-AAD-CMK and Ac-IETD-CHO, discussed below) also were
able to outcompete biotin-YVAD-CMK binding, whereas additional inhibitors that did not prevent activity (e.g., E-64 and
leupeptin) did not outcompete binding (data not shown). The
ligand-specific labeling of the 84-kD protein corresponds to the
substrate and inhibitor specificity identified for the saspase,
suggesting that the labeled 84-kD protein is the saspase. These
results were later confirmed during purification of the saspase
(see below).
Figure 3. The Caspase-Like Protease Is an 84-kD Ser Protease.
(A) Reversibility of inhibition of the protease in activity B. Three distinct
inhibitors all containing the VAD recognition motif were incubated at
a concentration of 200 mM with activity B for 2 h. Samples were washed
twice in buffer then reconstituted to original volume. After initial incubation and after each wash, aliquots were removed and assayed for
Z-VAD-AFC hydrolytic activity. Irreversible inhibition of the protease
was achieved only with the CMK-based inhibitor.
(B) Biotin blot of the protease in activity B. Each lane contains a 30-mL
sample of activity B from the HIC column. Samples were preincubated
2 h with a nonbiotinylated inhibitor (lanes 2 and 3) then incubated 2 h
with biotin-YVAD-CMK (lanes 1 to 3). The caspase-like protease shows
specific binding to biotin-YVAD-CMK because binding is competitive
with Z-VAD-CMK, an inhibitor, but not Z-DEVD-CMK, which does not
inhibit activity. Only the portion of the blot containing the 84-kD
protein is shown.
inhibit both Cys and Ser proteases, whereas FMK (fluoromethylketone)-based inhibitors irreversibly inhibit Cys proteases
but reversibly inhibit Ser proteases. CHO (aldehyde)-based
inhibitors reversibly inhibit both Cys and Ser proteases. All three
of these inhibitors inhibited activity of the protease by at least
80% when incubated for 2 h (Figure 3A, initial column). However,
after two washes, inhibition was lost from the FMK- and CHObased inhibitors, indicating reversible inhibition (Figure 3A, 1st
wash and 2nd wash columns). Only the CMK-based inhibitor
irreversibly inhibited activity, indicating that the caspase-like
protease is a Ser, not a Cys, protease. These results were later
confirmed by amino acid sequencing (discussed below). Because the protease(s) with caspase-like hydrolytic activity in
activity B is a Ser protease, we referred to this enzyme(s) as
a saspase.
Extracellular Localization of the Saspase after
Victorin Treatment
Infiltration of victorin into unpeeled leaves caused the release of
the saspase into the ECF. The use of leaves with the epidermis
removed provides a very sensitive and rapid assay of victorin
activity (Figure 1). However, because this assay cannot be used
for the collection of the ECF, a different assay was developed. In
this new assay, victorin was directly infiltrated into whole leaf
segments, thus allowing the ECF to be removed by centrifugation. However, this leaf infiltration assay is less sensitive. Infiltration of victorin at 1 mg/mL resulted in the first observation of
Rubisco proteolysis and DNA laddering (discussed below) at 8 h
after infiltration. Observation of levels of Rubisco proteolysis and
DNA laddering comparable to that seen after 4-h incubation of
peeled A. sativa leaf segments in 20 ng/mL of victorin were not
seen until 15 h after infiltration (data not shown). Thus, the relative
rates of Rubisco proteolysis and DNA laddering and, presumably, release of the saspases into the ECF are different in
the two assays.
ECF was collected from victorin-infiltrated leaves at 0 to 6 h
after infiltration with victorin, and then saspase activity was
measured by Z-VAD-AFC hydrolysis (Figure 4A). Protease
activity was first detected at 1 h and was maximal at 5 h. The
same ECF samples were incubated with 200 mM biotin-YVADCMK for 2 h, separated by SDS-PAGE, and blotted for biotin
visualization. The appearance of the 84-kD saspase band in the
ECF (Figure 4B) coincided with hydrolytic activity. Four-centimeter leaf sections were used for extraction of the ECF, and
typically, eight leaf sections were used per treatment/experiment. For the first 5 h (the time when maximum saspase activity
Caspase-Like Ser Proteases
861
had no effect on accumulation of the saspase in the ECF (data
not shown).
Heat Shock–Induced PCD Is Similar to
Victorin-Induced PCD
Figure 4. Release of the Saspase into the ECF after Victorin Treatment.
(A) and (B) Leaves were infiltrated with 1 mg/mL of victorin (vic) or water
(no victorin control), and the ECF was removed at the indicated time.
(A) Z-VAD-AFC hydrolytic activity in the ECF. Activity expressed as
relative fluorescence units.
(B) Biotin blot of the ECF. Twenty microliters of each sample was incubated with 200 mM biotin-YVAD-CMK for 2 h before electrophoresis and
blotting.
(C) and (D) Leaves were preinfiltrated with 400 mM biotin-YVAD-CMK,
preincubated 2 h, then infiltrated with 1 mg/mL of victorin mixed with
100 mM biotin-YVAD-CMK.
(C) Biotin blot of the ECF.
(D) Biotin blot of total protein extract. Note that the saspase is uniformly
labeled in all samples including the water only/no victorin control (last
lane).
was recovered), the quantity of ECF recovered in untreated
leaves ranged from 6.5 to 8 mL/leaf segment and averaged 7.5 mL
(n ¼ 56), whereas toxin-treated leaf segments ranged from 6.5 to
10.75 mL/leaf segment and averaged 9 mL (n ¼ 52). Slightly more
ECF was consistently recovered from toxin-treated leaf segments at all time points likely because of the fact that toxin
causes substantial leakage of electrolytes (e.g., Wheeler and
Black, 1962; Sammadar and Scheffer, 1968), which would
increase the water potential of treated cells.
Biotin-YVAD-CMK prevented Rubisco proteolysis with the
same effectiveness as the other caspase inhibitors (data not
shown), and furthermore, infiltration of leaves with biotin-YVADCMK provided in vivo labeling of the saspase. In Figure 4C, the
ECF was collected from leaves that were preinfiltrated with 400
mM biotin-YVAD-CMK for 2 h and then infiltrated with 1 mg/mL of
victorin mixed with 100 mM biotin-YVAD-CMK. Accumulation of
the saspase in the ECF (Figure 4C) followed approximately the
same pattern as shown in Figure 4B (activity could not be
measured because of inhibition by biotin-YVAD-CMK). Analysis
of total cellular protein prelabeled with biotin-YVAD-CMK (Figure
4D) indicates a constant level of saspase labeling throughout all
time points, including the 6 h no victorin control (Figure 4D, last
lane). These results suggest that transcriptional or translational
activation of the saspase is apparently not required. In addition,
treatment of leaves with cyclohexamide before victorin treatment
Heat shock has been successfully used to induce PCD in
Cucumis sativus (cucumber) (Balk et al., 1999) and N. tabacum
(Tian et al., 2000). We wanted to ascertain weather or not heat
shock could induce characteristics similar to victorin-induced
PCD in A. sativa. Victorin-sensitive and -insensitive leaves were
infiltrated with water (Figures 5A to 5D, lanes 1 and 4), 1000 ng/
mL of victorin (Figures 5A to 5D, lanes 2 and 5), or heat shock
(458C for 1 h) (Figures 5A to 5D, lanes 3 and 6). The ECF was
collected 4 h after start of the treatment, and protein and DNA
samples were collected after 15 h. Heat shock induces the
typical characteristics of victorin-induced PCD in victorin-sensitive and -insensitive A. sativa, including Rubisco proteolysis
(Figure 5A, lanes 3 and 6), DNA laddering (Figure 5B, lanes 3 and
6), and release of the 84-kD saspase into the ECF (Figures 5C and
5D, lanes 3 and 6). The water controls for both sensitive (Figure 5,
lane 1) and insensitive (Figure 5, lane 4) leaves did not display any
of the characteristics associated with PCD, nor did victorintreated insensitive leaves (Figure 5, lane 5). Heat shock–induced
Figure 5. Heat Shock–Induced PCD in Victorin-Sensitive and -Insensitive A. sativa.
Victorin-sensitive and -insensitive A. sativa leaves were infiltrated with
water (lanes 1, 3, 4, and 6) or 1 mg/mL of victorin (lanes 2 and 5). Waterinfiltrated leaves in lanes 3 and 6 were heat shocked for 60 min at 458C
and then incubated at 258C for indicated times below.
(A) Coomassie blue–stained SDS-polyacrylamide gel of total protein
extracted 15 h after treatment. Rubisco cleavage is seen in sensitive
leaves treated with victorin and sensitive and insensitive leaves that were
heat shocked. Only the region containing Rubisco is shown.
(B) 1.5% agarose gel of DNA extracted 15 h after treatment.
(C) Biotin blot of ECF collected 4 h after treatment. Samples were
incubated with 200 mM biotin-YVAD-CMK for 2 h before electrophoresis.
(D) Z-VAD-AFC hydrolytic activity in the ECF. Activity expressed as
relative fluorescence units.
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Figure 6. Inhibition of Heat Shock–Induced Rubisco Proteolysis.
A. sativa leaves were preincubated with 400 mM of indicated inhibitor for
2 h and then heat shocked for 60 min at 458C (sample in lane 1 did not
undergo heat shock). Proteins were isolated 15 h after heat shock. Only
the region containing Rubisco is shown.
phy. SAS-1 and SAS-2 possessed nearly identical NH2-terminal
sequences, with the only difference being the lack of the first Thr
residue at the NH2 terminus of SAS-1 (Table 2). Consistent with
the inhibitor studies implicating a Ser protease, the protein
sequence of the NH2 terminus of both SAS-1 and SAS-2 were
homologous to the mature NH2 terminus of several plant
subtilisin-like Ser proteases (Table 2). Additional sequence information was obtained from three internal peptides produced
by a partial tryptic digest of SAS-2 (no internal sequence data
was obtained from SAS-1). These three peptides also exhibited
homology to plant subtilisin-like Ser proteases (Table 2). A
putative subtilisin-like Ser protease from rice (Oryza sativa) was
the most similar protein identified (77.4%; 48 out of 62 amino
acids identical).
Characterization and Comparison of SAS-1 and SAS-2
Rubisco proteolysis, like that induced by victorin, was prevented
by the caspase inhibitors Ac-VAD-CMK and Ac-DEVD-CMK, the
granzyme B inhibitor Ac-AAD-CMK, and the general Cys
protease inhibitors E-64 and leupeptin (Figure 6).
Purification of Saspase-1 and Saspase-2
The purification protocol from 200 g of victorin-treated (100 ng/
mL) A. sativa leaves is summarized in Table 1. The proteolytic
activity in the crude extract and the 50% ammonium sulfate
fraction was either too dilute to obtain an accurate measurement
of activity or contained an inhibitory compound because total
activity (Table 1, column 3) more than doubled after the HIC
column. Anion exchange chromatography separated the saspase activity into two closely eluting peaks (Figure 7): the first
was termed Saspase-1 (SAS-1) and the second Saspase-2
(SAS-2). SAS-1 and SAS-2 were purified to homogeneity by size
exclusion chromatography and both migrated in an SDS-PAGE
with an apparent mass of 84 kD, as detected by silver staining
(Figure 8).
SAS-1 and SAS-2 Are Subtilisin-Like Ser Proteases
NH2-terminal amino acid sequencing was performed on gelpurified SAS-1 and SAS-2 after anion exchange chromatogra-
Matrix-assisted laser-desorption ionization time of flight (MALDITOF) mass spectrometry performed on purified SAS-1 and
SAS-2 indicated that both masses were 74,500 D 6 1900 D
(Figure 9). Because of the size of the protein analyzed and the
sensitivity of the instrument, the peak width was too large to
obtain a more accurate measurement of mass. Nevertheless,
these data indicate that the actual molecular mass of both SAS-1
and SAS-2 is 10,000 D smaller than that indicated by their
migration in SDS-PAGE (cf. Figures 7 and 8 with Figure 9).
Peptide maps were created from MALDI-TOF mass spectrometry analysis of a partial tryptic digest of both SAS-1 and
SAS-2 (Figure 10). A comparison of the two peptide maps
indicate that SAS-1 and SAS-2 have in common all major peaks
in the profile, and furthermore, the mass of the peaks are nearly
identical. Two major peaks with mass differences are indicated
with arrows in Figure 10. In both cases, the mass of the peak is
larger in SAS-1, with peak A having a mass difference of 16.1 D
and peak B of 14.21 D. This size variation may indicate a single
amino acid substitution or a difference in posttranslational
modifications.
The effect of pH on hydrolytic activity was assayed for SAS-1
and SAS-2 with Z-VAD-AFC as a substrate. Figure 11 illustrates
that maximal activity for both enzymes is observed at pH 6.5, and
SAS-1 and SAS-2 also share similar patterns of activity as the pH
is increased or decreased.
Table 1. Purification of SAS-1 and SAS-2 from 200 g of Victorin-Treated Leaves
Purification Step
Crude extract
(NH4)2SO4 pellet
HIC column
Heparin affinity
Anion exchange
SAS-1
SAS-2
Size exclusion
SAS-1
SAS-2
a1
Total Protein
(mg)
Total Activity
(pmol/min)a
812
380
45.46
12
1577.6
1520.0
3957.4
2502.4
1.9
4.0
87.1
208.5
1
2.1
44.8
107.4
0.082
0.106
844.3
1087.7
10296.3
10261.3
5307.4
5289.3
53.5
68.9
0.042
0.054
573.9
739.4
13664.3
13692.6
7043.5
7058.1
36.4
46.9
pmol of substrate cleavage ¼ 6.024 fluorescence units.
Specific Activity
(pmol/min/mg)a
Purification
Fold
Yield (%)
100
96
250
158
Caspase-Like Ser Proteases
863
Figure 7. Anion Exchange Chromatography Separated Two Distinct Peaks of Saspase Activity.
Silver stained gel of active fractions (20 mL loaded/lane) eluted off the anion exchange column. Z-VAD-AFC hydrolytic activity of each sample is
indicated below the gel, expressed as relative fluorescence units. Fractions 42 to 52 (SAS-1) and 54 to 64 (SAS-2) were pooled for individual purification
of SAS-1 and SAS-2 by size exclusion chromatography.
Salts generally enhanced saspase activity. The effects of
different salts on Z-VAD-AFC hydrolysis by SAS-1 and SAS-2 are
presented in Table 3. Monovalent salts (i.e., NaCl and KCl) were
less efficient at stimulating hydrolysis than divalent salts (i.e.,
MgCl2 and CaCl2). The heavy metal salt ZnCl2 inhibited activity
with or without the addition of NaCl.
SAS-1 and SAS-2 were tested for hydrolytic activity against
a variety of caspase and granzyme B–specific peptide substrates (Table 4). The two enzymes displayed indistinguishable
substrate recognition specificity, whereas SAS-2 possessed
slightly higher hydrolysis rates when standardized against
Z-VAD-AFC hydrolysis. Furthermore, neither saspase hydrolyzed any non-caspase substrate tested (with the exception of
the granzyme B substrate), including two subtilisin A substrates
and the general protease substrate casein (Table 5), nor did they
cleave purified Rubisco or BSA (data not shown). These data
indicate that both SAS-1 and SAS-2, like caspases, specifically
cleave after Asp residues and, also similar to caspases, possess
specificity beyond recognition of the P1 residue because not all
substrates with Asp in the P1 position are hydrolyzed by SAS-1
or SAS-2.
The effects of various caspase and the granzyme B inhibitors
on SAS-1 and SAS-2 Z-VAD-AFC hydrolytic activity are
summarized in Table 6. Similar to their substrate recognition
profiles, SAS-1 and SAS-2 also share nearly identical inhibitor
profiles. Several non-caspase-specific protease inhibitors were
tested for their ability to prevent Z-VAD-AFC hydrolysis (Table 7).
Only the subtilisin-specific inhibitor Z-GF-NHO-Bz had a strong
inhibitory effect on both SAS-1 and SAS-2, whereas all other
inhibitors, including another subtilisin-specific inhibitor, BocAPF-NHO-Bz, had only an intermediate or weak inhibitory effect
on hydrolytic activity of both enzymes.
Autoproteolysis of SAS-1 and SAS-2
Gel-purified (post-anion exchange chromatogaphy) SAS-1 and
SAS-2 were used to produce polyclonal antibodies in rabbits.
The antisera produced from SAS-1 and SAS-2 were crossreactive with each other (data not shown). Because SAS-2
antiserum had a higher titer of polyclonal antibodies, it was used
for protein gel blot analysis.
A protein gel blot of purified SAS-1 and SAS-2 probed with
SAS-2 antiserum is shown in Figure 12. After addition of SDSloading buffer to each sample (200 ng purified enzyme), the
samples were either boiled 5 min (Figure 12, lanes 2 and 4) or not
boiled (Figure 12, lanes 3 and 5) before loading on the gel.
Nonboiled samples of both enzymes underwent autoproteolysis
and accumulated lower molecular mass bands. The SAS-2
sample contained four distinct proteolytic bands—a, b, c, and
d— whereas the SAS-1 sample only contained three bands—a,
c, and d. Bands c and d are more visible when overexposed (data
not shown).
DISCUSSION
PCD and a Protease Cascade
Figure 8. Electrophoresis of SAS-1 and SAS-2 after Purification by Size
Exclusion Chromatography.
Silver stained gel of purified SAS-1 and SAS-2; 15 mL of active fraction
was loaded per lane.
The PCD response in plants is currently under intense characterization by many research laboratories. Apoptosis is the most
well-characterized form of PCD. Because caspases can play
such a critical role in the process and because numerous experiments have shown that caspase-specific inhibitors prevent
864
The Plant Cell
Table 2. Comparison of Partial Amino Acid Sequence of SAS-1 and SAS-2 with Other Known Subtilisin-Like Ser Proteases
Protein
Amino Terminus
Internal Peptide 1
SAS-1
SAS-2
Rice subtilisin
P69A
P69B
P69C
ARA12
Cucumisin
-THTPEFLGLSAAGG-LWEAS-EYG
TTHTPEFLGLSAAGG-LWEAS-EYG
109-TTHTPEFLGVSGAGG-LWETA-SYG-133
115-TTHTSSFLGLQQNMG-VWKDS-NYG-137
115-TTHTPSFLGLQQNMG-VWKDS-NYG-137
115-TTHTPSFLGLQQNMG-LWKDS-NYG-137
107-TTRTPLFLGLDEHTADLFPEAGSYS-131
111-TTRSWDFLGFPLTVP---RRS-QVE-131
----------VHPDWSPAAVR
545-VHPEWSPAAIR-556
548-THPDWSPAAIK-558
547-THPDWSPAVIK-557
547-SHPDWSPAVIK-557
558-VHPEWSPAAIR-568
541-YNPTWSPAAIK-551
Protein
Internal Peptide 2
Internal Peptide 3
SAS-1
SAS-2
Rice subtilisin
P69A
P69B
P69C
ARA12
Cucumisin
--------------EVTNVGDGPASYTAK
674-VVTNVGAGAASYRAK-688
667-TVTNVGDAKSSYKVE-681
666-TVTNVGDATSSYKVE-680
667-TVTNVGDAKSSYTVQ-681
666-NVDGVGAYKYTRTVT-679
655-TLTSVAPQASTYRAM-670
------------SPIVATTASSTPF
748-SPIVATTLSSTRL-760
737-SPI-ALLLIQ----745
738-SPI-AVV-SA----745
739-SPI-AVEFALATK-750
750-SP-VAISWT-----757
723-SPITITSLV-----731
Rice subtilisin, putative subtilisin-like protease (accession number BAB89803). P69A, P69B, and P69C, subtilisin-like proteases from L. esculentum
(accession numbers CAA76724, CAA76725, and T06577, respectively). ARA12, subtilisin-like protease from Arabidopsis (accession number
NP_569048). Cucumisin, subtilisin-like protease form Cucumis melo (accession number A55800). Identical amino acids are in bold.
some forms of PCD in plants (del Pozo and Lam, 1998; Sun et al.,
1999; Tian et al., 2000; Richael et al., 2001; Elbaz et al., 2002;
Mlejnek and Procházka, 2002), it has been speculated that
proteases similar to caspases are involved in plant PCD. We
have purified and characterized two subtilisin-like Ser proteases
that possess specificity for caspase-specific substrates and
inhibitors and apparently participate in a signaling cascade
involving multiple proteases, leading to the proteolysis of
Rubisco. The proteases have been referred to as saspases.
Figure 9. Molecular Mass of both SAS-1 and SAS-2 Is 74.5 kD as
Indicated by Mass Spectrometry.
Spectrograph of purified SAS-1 (top spectrograph) and SAS-2 (bottom
spectrograph) were individually analyzed by MOLDI-TOF mass spectrometry. First peak is the doubly charged ion, and the second peak is the
singly charged ion.
The saspase proteases were originally implicated based on
the effects of inhibitors that prevented victorin and heat shock–
induced Rubisco proteolysis, a characteristic of PCD in A. sativa.
Other inhibitors that prevent this response (Ac-DEVD-CMK,
E-64, and leupeptin) do not inhibit the activity of either saspase,
indicating the involvement of additional proteases. The identification of another protease with specificity for DEVD derived
substrates and inhibitors, but not E-64 or leupeptin, suggested
that at least two other proteases function within a protease
cascade, with either or both of the saspase enzymes, leading to
the proteolysis of Rubisco.
This evidence suggests that the saspases are components of
a PCD-induced protease cascade that ultimately leads to the
activation of a protease that is targeted to the chloroplast to
cleave Rubisco. Although cleavage of Rubisco is inhibited by AcVAD-CMK and Z-AAD-CMK, both of which are recognized by the
saspases, it is not likely that either saspase directly cleaves
Rubisco in vivo. Four lines of evidence suggest that an alternative
protease cleaves Rubisco: (1) E-64, leupeptin, and Ac-DEVDCMK prevent Rubisco cleavage but do not affect SAS-1 or
SAS-2 activity or their release into the ECF; (2) the cleavage site in
Rubisco (Navarre and Wolpert, 1999) does not conform to the
substrate specificity for the saspases; (3) purified SAS-1 and
SAS-2 do not cleave Rubisco in vitro (data not shown); and (4) the
saspases appear to function extracellularly.
A Gene Family
The caspase-like activity was purified into two distinct proteases, SAS-1 and SAS-2. The two proteases are nearly
indistinguishable, in that their substrate and inhibitor profiles
are identical, they have the same pH and salt requirements, they
are predicted to have equivalent molecular masses determined
Caspase-Like Ser Proteases
865
been annotated as being subtilisin-like Ser proteases. Rice has
had only one subtilisin-like Ser protease gene characterized
(Yamagata et al., 2000), but seven have been annotated
from the genome, two of which have high sequence similarity
to SAS-1 and SAS-2. Thus, it is likely that two or more subtilisinlike Ser proteases are present in A. sativa, and quite possibly a
more divergent protease with DEVD specificity functions along
with SAS-1 and SAS-2 within the protease cascade leading to
Rubisco proteolysis. Because of the structural and immunological similarities between the saspases and the difficulty in
purifying large amounts from the ECF, we have not been able
to distinguish which saspase, if not both, is released into the ECF.
However, initial studies have indicated that after induction of
PCD and removal of the ECF from victorin-treated leaves, total
saspase protein remaining in the whole leaf is dramatically
reduced, suggesting that both SAS-1 and SAS-2 are released
into the ECF. Further experiments need to be performed to
confirm these observations.
Processive Protease and Family Relations
Figure 10. Peptide Maps of SAS-1 and SAS-2.
Partial tryptic digest of SAS-1 and SAS-2 analyzed by MALDI-TOF mass
spectrometry. Major peptide peaks with molecular mass differences
between SAS-1 and SAS-2 are indicated by an arrow. m/z, mass-tocharge ratio.
by either electrophoretic migration or mass spectrometry, they
possess similar peptide maps, they have an identical NH2terminal sequence (except the first amino acid of SAS-1), and
they both bind heparin and are autoproteolytic. Although they
recognize the same substrates and inhibitors, SAS-2 does have
a higher hydrolysis rate for several caspase substrates and is
slightly more sensitive to some caspase inhibitors. Furthermore,
the difference in mass between two of the several major peaks in
their peptide maps suggest a differential posttranslational
modification (such as a methyl group) or a single amino acid
substitution. The comparisons between SAS-1 and SAS-2
indicate that they are different yet highly similar proteases, most
likely from two separate genes. But the possibility exists that they
are differentiated by unique posttranslational modifications to
the same gene product.
Gene families of subtilisin-like Ser proteases have been
identified in several plant species. Lycopersicon esculentum
(tomato) has had 10 genes for subtilisin-like Ser proteases thus
far cloned and characterized (Vera and Conejero, 1988; Tornero
et al., 1997; Jordá et al., 1999, 2000; Meichtry et al., 1999; Janzik
et al., 2000; Riggs et al., 2001). Whereas Arabidopsis (Arabidopsis thaliana) has had only four genes characterized
(Neuteboom et al., 1999; Berger and Altmann, 2000; Tanaka
et al., 2001; Hamilton et al., 2003), at least 10 more genes have
The substrate specificity of SAS-1 and SAS-2 is distinct from all
other known subtilisin-like Ser proteases. Among the substrates
tested, only certain caspase substrates and the granzyme B
substrate were hydrolyzed and to varying degrees, indicating that
although an aspartate residue is required in the P1 position,
specificity extends beyond this one residue. No other identified
protease from plants has been reported to be specific for
caspase substrates or to cleave exclusively after an aspartate
residue. Plant subtilisin-like Ser proteases differ widely in their
substrate specificity. Cucumisin (Yamagata et al., 1994), macuralisin (Rudenskaya et al., 1995), taraxalism (Rudenskaya et al.,
1998), hordolisin (Terp et al., 2000), and Ser endopeptidase-1
(Fontanini and Jones, 2002) have been reported as having a broad
substrate range and are thought to be degradative proteases,
whereas Auxin Induced in Root3 (Neuteboom et al., 1999),
Abnormal Leaf Shape1 (Tanaka et al., 2001), Stomatal Density
Figure 11. Effects of pH on Hydrolytic Activity of SAS-1 and SAS-2.
Partially purified SAS-1 and SAS-2 (post-anion exchange chromatography) were assayed for hydrolytic activity at different pHs with the
substrate Z-VAD-AFC. RFU, relative fluorescence units.
866
The Plant Cell
Table 3. Effect of Different Salts on Hydrolytic Activity of SAS-1 and
SAS-2
Activity
Salt
Concentration
SAS-1
SAS-2
NaCl
10 mM
100 mM
500 mM
10 mM
100 mM
500 mM
10 mM
100 mM
10 mM
100 mM
10 mM
100 mM
10 mM
100 mM
15 6 1.6
48 6 7.1
95 6 2.7
13 6 3.3
42 6 0.8
108 6 6.1
53 6 3.1
102 6 4.3
50 6 1.9
90 6 2.2
16 6 1.5
20 6 4.2
25 6 6.1
1 6 0.5
11 6 3.1
41 6 0.8
100 6 0.4
6 6 5.0
51 6 3.4
128 6 10.7
65 6 9.8
118 6 3.9
65 6 2.1
113 6 3.7
28 6 2.8
18 6 5.6
32 6 2.7
1 6 1.0
KCl
CaCl2
MgCl2
ZnCl2
ZnCl2 (with 500 mM
NaCl)
Partially purified enzymes assayed 1 h in indicated salt with the substrate Z-VAD-AFC. Activity is expressed as relative fluorescence units.
and Distribution1 (SDD1) (Berger and Altmann, 2000), Tomato
subtilase1 (LeSBT1) (Janzik et al., 2000), 50-kD Systemin Binding
Protein (Schaller and Ryan, 1994), and Ara12 (Hamilton et al.,
2003) are described as possessing greater substrate specificity
and are thought to function as processive proteases. Until more is
known about the biochemical and physiological properties of
plant subtilisin-like Ser proteases, the biological significance of
the difference between a degradative or processive function may
prove challenging to determine. Because SAS-1 and SAS-2 do
not generally degrade proteins or non-caspase-specific substrates (e.g., casein), require an aspartate residue in the P1
position of the substrate, and are likely involved in a signaling
cascade, it appears evident that they function as processive
proteases.
Plant subtilisin-like Ser proteases are members of the
Pyrolysin family of subtilisin-like Ser proteases (Siezen and
Leunissen, 1997) and are predicted, based on DNA sequence of
the cloned proteases, to be expressed as pre-pro-protein
precursors. Another family of subtilisin-like Ser proteases, the
Kexin family, also is expressed as pre-pro-protein precursors.
Kex2 from Saccharomyces cerevisiae, the prototype member of
this family, has been well-characterized (Mizuno et al., 1988). In
Kex2, the pre-domain is a signal peptide that directs the enzyme
into the secretory system of the endoplasmic reticulum (ER) and
is cleaved during import (Van de Ven et al., 1991). The prodomain is autocatalytically cleaved while in the ER, which results
in the formation of the active, mature form of the protease
(Gluschankof and Fuller, 1994). The pre- and pro-domains of
plant subtilisin-like Ser proteases are thought to each function in
a similar manner to those of the Kex2 enzyme because several
plant subtilisin-like Ser proteases have been localized extracellularly (Vera and Conejero, 1988; Yamagata et al., 1994; Taylor
et al., 1997), and the NH2 terminus of mature, active enzymes
corresponds to the predicted mature NH2 terminus absent the
pre- and pro-domains (Terp et al., 2000; Popovič et al., 2002).
The saspases from A. sativa are extracellularly localized during
the PCD response, and their NH2-terminal sequence is similar to
the mature, processed NH2 terminus of most plant subtilisin-like
Ser proteases, characteristics which correspond in function to
the pre-pro-protein model of subtilisin-like Ser proteases.
Furthermore, because pro-domains are autocatalytically processed and the pro-domain cleavage site indicates, to some
degree, the cleavage specificity for the enzyme, it might be expected that in the saspases, the amino acid preceeding the
cleavage site would be an aspartate. Interestingly, the protein
with the most similarity to the sequenced saspase peptides, the
putative protease annotated from the rice genome, does have an
aspartate residue in the P1 position of the predicted pro-domain
cleavage site, suggesting that the putative rice protease may
possess similar cleavage specificity to the saspases from
A. sativa.
Heparin and Autoproteolysis
In the early stages of characterization and before purification and
partial sequence analysis, it was clear that the saspases were
high molecular mass, extracellular Ser proteases apparently
involved in a protease cascade. That depiction strongly resembled a Drosophila melanogaster protease called gastrulation defective (GD), which is a 72-kD extracellular Ser protease
involved in a protease cascade leading to dorsal-ventral signaling (Konrad et al., 1998; Han et al., 2000; DeLotto, 2001;
Dissing et al., 2001; LeMosy et al., 2001). After purification and
partial sequencing, it became clear that GD and SAS-1 and
SAS-2 are structurally unrelated, although they may share some
similar biological characteristics. Two other characteristics were
later identified as being in common between the saspases and
GD, namely heparin binding and autoproteolysis.
Heparin, a sulfated glycosaminoglycan, is involved in several
biological activities in animals, including binding antithrombin III.
Table 4. Hydrolytic Activity of SAS-1 and SAS-2 toward Caspase and
Granzyme B Substrates
Substrate
Z-VAD-AFCa
Z-YVAD-AFCa
Z-VDVAD-AFCa
Z-DEVD-AFCa
Ac-LEVD-AFCa
Z-WEHD-AFCa
Ac-VEHD-AFCa
Ac-VEID-AFCa
Ac-VKMD-AFCa
Ac-VNLD-AFCa
Ac-IETD-AFCa
Ac-LEHD-AFCa
Z-AAD-AFCa
Protease with
Optimal
Specificity
Pan-caspase
Caspase-1
Caspase-2
Caspase-3
Caspase-4
Caspase-5
Caspase-6
Caspase-6
Caspase-6
Caspase-6
Caspase-8
Caspase-9
Granzyme B
Percentage of Z-VAD-AFC
SAS-1
SAS-2
100% 6 1.2%
19% 6 2.1%
0% 6 0%
0% 6 0%
39% 6 4.4%
0% 6 0%
311% 6 21.5%
0% 6 0%
623% 6 32.5%
384% 6 18.2%
130% 6 10.1%
105% 6 9.6%
144% 6 13.9%
100%
28%
0%
0%
51%
0%
366%
0%
654%
521%
172%
140%
183%
6 1.9%
6 2.5%
6 0%
6 0%
6 2.6%
6 0%
6 15.6%
6 0%
6 28.8%
6 21.8%
6 14.7%
6 10.2%
6 20.1%
Z-VAD-AFC was used as the 100% control.
sequence in single-letter amino acid code.
Z, benzyloxycarbonyl; Ac, acetyl; AFC, 7-amido-4-(trifluoromethyl)coumarin.
a Recognition
Caspase-Like Ser Proteases
867
Table 5. Hydrolytic Activity of SAS-1 and SAS-2 toward Various Protease Substrates
Percentage of Z-VAD-AFC
Substrate
Protease with Optimal Specificity
SAS-1
SAS-2
Z-VAD-AFCa
Casein
Boc-GGL-pNAa
A-AAL-pNAa
SY-AFCa
GF-AFCa
PR-AFCa
GP-AFCa
Z-RR-AMCa
Suc-AAPF-AMCa
Z-AKR-AMCa
H-L-AMCa
Pan-caspase
General protease
Subtilisin A
Subtilisin A
Dipeptydylpeptidase I
Dipeptydylpeptidase I
Dipeptydylpeptidase I
Dipeptydylpeptidase IV
Cathepsin B
Chymotrypsin
ICRM Ser protease I
Aminopeptidase
100% 6 2.8%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
100% 6 2.1%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
0% 6 0%
Z-VAD-AFC was used as the 100% control.
sequence in single-letter amino acid code.
Z, benzyloxycarbonyl; Boc, tert-butyloxycarbonyl; Suc, N-succinyl; AFC, 7-amido-4-(trifluoromethyl)coumarin; AMC, 7-amido-4-methylcoumarin.
a Recognition
Such binding accelerates the rate of antithrombin III–mediated
inhibition of Ser proteases involved in the blood coagulation
cascade (reviewed in Petitou et al., 1988; Capila and Linhardt,
2002). Heparan sulfate, a heparin derivative, is ubiquitously
distributed on the surfaces of animal cells and in the
extracellular matrix where it mediates various physiological
and pathological processes (reviewed in Capila and Linhardt,
2002). Though the biological significance of GD binding heparin
is not known, Dissing et al. (2001) suggest that the pipe gene
product, a heparan sulfate 2-O-sulfotransferase, might regulate
activity of GD through binding of a yet unknown sulfated
proteoglycan. Possibly, the binding of SAS-1 and SAS-2 to
a heparin-like molecule in plants may have a regulatory role,
similar to Ser proteases involved in the blood coagulation
cascade or the putative role of heparin-like molecules in
regulating GD.
Autoproteolysis by GD, SAS-1, and SAS-2 is an intriguing yet
confounding characteristic. GD has been shown to undergo
autoproteolysis only in the presence of the Snake protease and
into two distinct bands of 44 and 46 kD (Dissing et al., 2001).
Neither proteolytic activity nor a biological function has been
identified for the products of proteolysis, though it was
suggested that the unprocessed form of the protease was the
zymogen and the proteolytic products might function as the
active component of the cascade (Dissing et al., 2001). SAS-1
and SAS-2 autoproteolysis has only been observed in gel where
it is likely dependent on concentration and the state of
denaturation. Both proteases are stable at room temperature in
Table 6. Effects of Caspase and Granzyme B Inhibitors on SAS-1 and SAS-2 Hydrolytic Activity
Percent Activity of Noninhibited
Inhibitor
Preferred Protease Target
SAS-1
No inhibitor
Ac-VAD-CMKa
B-YVAD-CMKa
Ac-WEHD-CHOa
Ac-LDESD-CHOa
Ac-VDVAD-CHOa
Ac-DEVD-CMKa
Ac-DMQD-CHOa
Ac-LEVD-CHOa
Ac-VEID-CHOa
Ac-IETD-CHOa
Ac-LEHD-CMKa
Ac-LEED-CHOa
Z-AAD-CMKa
Pan-Caspase
Caspase-1
Caspase-1
Caspase-2
Caspase-2
Caspase-3
Caspase-3
Caspase-4
Caspase-6
Caspase-8
Caspase-9
Caspase-13
Granzyme B
100%
1%
1%
1%
39%
1%
90%
1%
1%
1%
1%
47%
68%
1%
SAS-2
6 3.0%
6 0.6%
6 0.1%
6 0.5%
6 2.4%
6 0.9%
6 4.9%
6 0.7%
6 0.2%
6 0.2%
6 0.4%
6 1.9%
6 3.2%
6 0.6%
100% 6 2.6%
1% 6 0.1%
1% 6 0.0%
1% 6 1.2%
49% 6 4.4%
2% 6 0.8%
91% 6 5.6%
1% 6 0.4%
1% 6 0.9%
1% 6 0.8%
1% 6 1.5%
58% 6 5.2%
67% 6 4.7%
1% 6 1.0%
Enzymes were preincubated with 200 mM inhibitor for 2 h and then assayed for Z-VAD-AFC hydrolytic activity for 1 h.
sequence in single-letter amino acid code.
Z, benzyloxycarbonyl; Ac, acetyl; B, biotin.
a Recognition
868
The Plant Cell
Table 7. Effects of Various Protease Inhibitors on SAS-1 and SAS-2
Hydrolytic Activity
Inhibitor
Preferred
Protease
Target
No inhibitor
Z-GF-NHO-Bza
Boc-APF-NHO-Bza
Aprotinin
AEBSFb
PMSFc
Leupeptin
Antipain
E-64d
PCMBe
Pepstatin A
Subtilisin
Subtilisin
Ser
Ser
Ser
Ser and Cys
Ser and Cys
Cys
Thiol
Aspartate
Percent Activity of Noninhibited
SAS-1
SAS-2
100% 6 2.2%
1% 6 0.5%
93% 6 4.2%
73% 6 8.1%
64% 6 5.4%
60% 6 5.9%
88% 6 2.5%
100% 6 3.9%
79% 6 5.1%
89% 6 4.0%
91% 6 2.5%
100% 6 3.0%
1% 6 2.2%
92% 6 3.8%
77% 6 2.8%
73% 6 4.2%
70% 6 3.4%
81% 6 2.5%
100% 6 2.7%
81% 6 3.2%
100% 6 1.9%
89% 6 4.1%
Purified SAS-1 and SAS-2 were preincubated with 200 mM inhibitor for
2 h, except PMSF (1 mM) and aprotinin (1 mg/mL), and then assayed for
Z-VAD-AFC hydrolytic activity for 1 h.
a Recognition sequence in single-letter amino acid code. NHO, nitrosyl;
Bz, benzoyl.
b 4-(2-aminoethyl)-benzenesulfonyl-fluoride.
c Phenylmethylsulfonyl fluoride.
d Trans-epoxysuccinyl-L-leucylamido-(4-guanidino)-butane.
e p-Chloromercuribenzoic acid.
biological buffers for several days (data not shown). Autoproteolysis by SAS-1 and SAS-2 is not unique among plant
subtilisin-like Ser proteases because two other proteases have
been shown to undergo a form of self-catalyzed proteolysis.
LeSBT1, a 73-kD protease from L. esculentum, is autoprocessed
into a 68-kD protease at pH 5.0 and 6, whereas it completely
autodegrades at pH 4.0 (Janzik et al., 2000), and hordolisin from
Hordeum vulgare (barley) was autoprocessed from 74 to 62 kD
during storage at 208C (Terp et al., 2000).
laddering, start to occur. This indicates that saspase appearance
in the ECF is not because of cell lysis but is a regulated event. This
conclusion is further supported by time-course studies (Curtis
and Wolpert, 2004) that have established that the plasma
membrane and tonoplast of victorin-treated A. sativa cells remain
structurally intact after the point at which Rubisco proteolysis,
DNA laddering, and even cell collapse occur.
The question arises as to where the saspases are localized
before release into the ECF. They could be retained in the ER
where they await processing and secretion. The Kexin family of
subtilisin-like Ser proteases is known to autocatalytically remove
their pro-domain in the ER before continuing in the secretory
pathway (Gluschankof and Fuller, 1994; Hill et al., 1995). A
mutation in the Kex2 protease that inhibits removal of the prodomain stimulates accumulation of proKex2 in the ER
(Gluschankof and Fuller, 1994). The pro-domain is thought to
contain an ER retention sequence. If the saspases are stored in
the ER while awaiting processing and secretion, then an inactive pro-saspase form should be identifiable. However, analysis
of proteins labeled in vivo with biotin-YVAD-CMK and extracted
under denaturing conditions (phenol extracted) from victorintreated and untreated tissue revealed similar concentrations of
only the mature processed proteases (Figure 4D). Soluble protein
extracts from victorin-treated and untreated tissue also contained comparable amounts of saspase activity (data not
shown). These data suggest that the saspases are likely
sequestered as active proteases.
Another possible saspase-retention area is the cell exterior,
where it could associate with the extracellular matrix or a membrane protein. Heparin or heparin-like molecules are typically
extracellular, either membrane bound or soluble (Petitou et al.,
1988; Stringer et al., 2002), and because the saspases are
ultimately extracellular, it is possible that a heparin-like molecule
Regulation and Localization
A unique characteristic of the saspases is that they appear to be
constitutively present in A. sativa cells, and release of active
saspase into the ECF but not de novo synthesis of the enzyme is
correlated with the induction of PCD. This is in contrast with other
proteases associated with plant PCD (Delorme et al., 2000;
Eason et al., 2002; Wan et al., 2002) or the HR (Vera and Conejero,
1988; Krüger et al., 2002), which are typically regulated at the
level of expression. P-69B and P-69C, members of a family of L.
esculentum subtilisin-like Ser proteases, are two pathogenesisrelated (PR) proteins that show increased expression levels
during HR (Jordá et al., 1999). The pathogen-induced P-69
enzymes also are secreted into the extracellular space (Tornero
et al., 1996a). However, unlike saspases, these proteins are not
present in the cell before induction of the HR (Tornero et al., 1997).
Release of the saspase(s) into the ECF occurs before the onset
of other markers of PCD, indicating that saspase release could be
causal to but is not a consequence of PCD. The saspase(s) starts
to appear in the ECF at 1 h after victorin treatment, at least 7 h
before other signs of PCD, such as Rubisco proteolysis and DNA
Figure 12. Autoproteolysis of SAS-1 and SAS-2.
Protein gel blot of purified SAS-1 and SAS-2 (200 ng each lane) boiled
and not boiled before electrophoresis. Nonboiled enzymes underwent
autoproteolysis with the appearance of lower molecular weight bands (a,
b, c, and d).
Caspase-Like Ser Proteases
plays a physiologically significant role, either before release of
the saspases or sometime during their biological function.
Whatever their storage location, the fact that the saspases
appear to be constitutively present in an active form suggests
that they can be rapidly released and that a quick response is
important for PCD.
PCD and Saspase Function
Though no physiological substrate(s) of the saspases has been
identified, data suggest that they activate, either directly or
indirectly, other proteases. Because the saspases recognize two
of the inhibitors (and their complementary substrates) that
prevent Rubisco proteolysis and because victorin treatment
affects localization of the saspases, it appears likely that they are
involved in the signal cascade leading to Rubisco proteolysis.
The saspases also likely function early in the signal cascade
because their release into the ECF is not prevented by any of the
protease inhibitors that prevent Rubisco proteolysis, indicating
that these inhibitors function either downstream or after release
of the saspases. Furthermore, because multiple proteases
appear to be involved in this cascade, one or both of the saspases are likely involved in activation of these other proteases.
Are these proteases the direct substrate, or do the saspases
interact with an extracellular protein, possibly membrane associated, which facilitates the activation of intracellular proteases?
Several reports have implicated the interaction of extracellular
proteases with Leu-rich repeat (LRR) proteins (LRP) during the
HR and developmental processes. The pathogen-induced,
subtilisin-like Ser protease P-69 from L. esculentum is able to
proteolytically process an extracellular LRP (Tornero et al.,
1996b). The LRP, predicted to be soluble in the extracellular
matrix, also has increased expression levels during pathogen
attack, but the function of the processed or unprocessed form is
yet to be determined. Krüger et al. (2002) describe an extracellular Cys protease, Required for Cladosporium fulvum Resistance3 (Rcr-3), that is required for Cf-2–dependent HR. Cf-2
is a resistance gene in L. esculentum that encodes a transmembrane LRP with an extracellular LRR domain. Not only is
Rcr-3 required for Cf-2–dependent HR, it also is necessary for
suppression of Cf-2–dependent autonecrosis. Two other extracellular proteases, SDD1, a subtilisin-like Ser protease (Berger
and Altmann, 2000), and Brassinosteroid-Insensitive1, a Ser
carboxypeptidase (Li et al., 2001), have been described as
necessary for stomata development and brassinosteroid signaling, respectively. Both proteases function upstream of LRR
receptor–like proteins that are required for continuation of their
respective signaling pathways (Li et al., 2001; Serna and Fenoll,
2002). Whether or not the saspases interact with another
protease, an LRR receptor–like protein, or a ligand for a membrane receptor is currently not known. Identification of the
physiological substrate for the saspases will be an important step
in furthering the understanding of the processes of PCD in
A. sativa.
Recently, Woltering et al. (2002) asked the question, ‘‘Do plant
caspases exist?’’ This research addresses that question by
providing insight into enzymes that possess caspase-like
activity, though not structurally related to caspases. Numerous
869
studies have indicated the involvement of caspase-like proteases in the regulation of plant PCD. Caspase-specific inhibitors
prevent PCD in response to bacteria (del Pozo and Lam, 1998;
Richael et al., 2001), fungal elicitors (Elbaz et al., 2002), and
various forms of stress (Sun et al., 1999; Tian et al., 2000; Elbaz
et al., 2002; Mlejnek and Procházka, 2002; Woltering et al., 2002).
Although these reports indicate that there are proteases in plants
that are inhibited by caspase-specific inhibitors and implicated in
PCD, no enzyme had yet been identified that pertains to these
observations. Saspases, plant enzymes possessing caspaselike specificity and associated with the PCD response, are
subtilisin-like Ser proteases, and partial amino acid sequence
analysis revealed homology to all plant subtilisin-like Ser
proteases. The unusual requirement for aspartate residues in
the P1 position of the substrate, the specificity of the enzyme
among aspartate-containing substrates, the lack of general
proteolytic activity, and the possible involvement in a signal
cascade suggest that the saspases function as processive
proteases. Release of one or both of the saspases into the ECF
occurs during victorin and heat shock–induced PCD, two
treatments that share other characteristics of PCD, namely
Rubisco proteolysis and DNA laddering. The saspases appear to
be involved in a PCD-induced signaling cascade based on
inhibitor studies and relocalization during the PCD response, but
unfortunately no direct evidence of their involvement has been
identified. Initial experiments to induce PCD in A. sativa leaves by
infiltration of purified saspase have provided interesting but
inconsistent results. The ability to purify large amounts of enzyme
(needed for these types of experiments) without first cloning the
gene has been the largest impediment. The identification of the
putative protease from rice, which possesses the highest amino
acid sequence similarity, should facilitate cloning of the saspase
genes from A. sativa. It also will be of interest to see if this putative
protease from rice has caspase-like activity and if it is involved in
rice PCD responses.
METHODS
Experimental Materials
A. sativa seedlings of the line X469 (victorin sensitive) and X424 (victorin
insensitive) were grown in a growth chamber for 6 to 7 d under a 16-h
photoperiod at 248C. All protease substrates and caspase inhibitors were
purchased from Calbiochem (La Jolla, CA). The remaining chemicals and
protease inhibitors were purchased from Sigma (St. Louis, MO).
Treatment of A. sativa Tissue
A. sativa leaves were treated with victorin by one of three methods. A.
sativa leaves, with their epidermis removed, were floated on 20 mM 3-(Nmorpholino)-propanesulfonic acid (Mops), pH 6.5, supplemented with 20
ng/mL of victorin. For leaf infiltration assays, intact A. sativa leaves were
infiltrated with 1 mg/mL of victorin in water and then cut into 4-cm leaf
segments. The third method, which was used to purify large quantities
of protein, treated leaf slices (2- to 3-mm cross-sectional pieces) with
100 ng/mL of victorin in water.
For heat shock treatments, 4-cm water-infiltrated leaf segments were
incubated for 60 min at 458C. Leaves were water infiltrated to facilitate
collection of ECF. After heat shock, leaves were incubated at 258C for the
times indicated before collection of the ECF, total protein, and DNA.
870
The Plant Cell
Total protein and DNA were extracted from leaf samples by
homogenizing 1 cm of leaf tissue in 200 mL of phenol combined with
200 mL of Tes buffer (50 mM Tris-HCl, pH 7.6, 5 mM EDTA, and 2% [w/v]
SDS). After homogenization, the phases were thoroughly mixed and
separated by centrifugation for 10 min. The protein (phenol) phase was
removed, and the proteins were precipitated by mixing with 53 volume
0.1 M ammonium acetate and 2.5% (v/v) b-mercaptoethanol in cold
methanol and incubating the mixture a minimum of 4 h at 48C. Samples
were then centrifuged for 10 min to pellet the proteins, washed in cold
methanol, and centrifuged again. Precipitated samples were dried and
resuspended in 100 mL of SDS-loading buffer (62.5 mM Tris-HCl, pH 6.8,
2.3% [w/v] SDS, 5% [v/v] b-mercaptoethanol, and bromophenol blue
[trace amount]). For SDS-PAGE analysis of Rubisco proteolysis, 8 mL of
sample was loaded on the gel. For biotin blots of total cellular protein,
30 mL of sample was loaded on the gel.
The DNA (aqueous) phase was precipitated by mixing with 2.53
volume cold ethanol and 0.13 volume 3 M Na acetate and incubating
overnight at 208C. DNA samples were centrifuged for 10 min, washed in
70% (v/v) cold ethanol, and centrifuged again. Precipitated DNA samples
were dried and resuspended in 20 mL of TE buffer (10 mM Tris-HCl, pH
7.5, and 1 mM EDTA) supplemented with 40 mg/mL of RNase. The entire
sample was loaded on a 1.5% (w/v) agarose gel.
ECF was collected by cutting the 4-cm leaf segments into two, 2-cm
segments and placing them vertically in a 1.5-mL microcentrifuge tube
with a pinhole in the bottom. The tubes were placed inside a collection
tube and centrifuged for 10 min at 1000g in a swinging bucket rotor. To
assay for proteolytic activity, 10 mL of ECF was used per reaction. For
biotin blots, 30 mL of biotin-YVAD-CMK–labeled ECF was loaded per lane
for each gel.
Protease Activity Assays
Substrate hydrolysis was measured, unless otherwise indicated, by
adding 10 mL of an enzyme sample to 90 mL of 20 mM Mops, pH 6.5, and
0.5 M NaCl supplemented with 20 mM of the indicated substrate for a final
volume of 100 mL. Samples were incubated at 258C for 60 min in 96-well
microtiter plates. Fluorescence produced by hydrolysis of the substrates
was read with a SpectroMax Gemini fluorometer (Molecular Devices;
Sunnyvale, CA). All assays were performed in triplicate.
Reversible and Irreversible Binding Experiments
Activity B from the HIC column was assayed for inhibition with three
inhibitors: Z-VAD-CHO, Z-VAD-FMK, and Ac-VAD-CMK. The inhibitors
(200 mM) were each added to 200 mL of sample equilibrated in 20 mM
Mops, pH 6.5, 1 mM DTT, and 100 mM NaCl, then incubated for 2 h. Tenmicroliter aliquots from each treatment were assayed for VAD-AFC
hydrolysis (see above). Samples were washed with 1 mL of 20 mM Mops,
pH 6.5, 1 mM DTT, and 100 mM NaCl and concentrated back to the
original volume with a 2-mL Centricon-30 (30,000 molecular weight cutoff
[MWCO]) (Amicon, Beverly, MA). Three, 10-mL aliquots were assayed
after each wash step.
Purification of SAS-1 and SAS-2
Soluble Protein Preparation and Ammonium Sulfate Precipitation
A. sativa leaves (200 g) were sliced into 2- to 3-mm pieces and incubated
4 h with 100 ng/mL of victorin in water. The slices were then homogenized
with a mortar and pestle in cold 20 mM Mops, pH 7.0, with 1 mM DTT. The
homogenate was filtered through four layers of cheesecloth and
centrifuged for 10 min at 10,000g. The resultant supernatant was removed
and centrifuged for 1 h at 100,000g. The soluble protein solution was
mixed with 50% (w/v) ammonium sulfate for 30 min at 48C, centrifuged for
30 min at 10,000g, and the pellet was solubilized in 20 mM Mops, pH 7,
containing 1 mM DTT and 1 M ammonium sulfate. The solubilized pellet
was mixed for 30 min at 48C, centrifuged for 10 min at 10,000g to remove
insoluble material, and filtered through a 0.22-mm filter.
HIC
Before the establishment of a purification protocol, initial characterization
of protease activity was performed on soluble protein extracted from 60 g
of A. sativa tissue (prepared as above but without ammonium sulfate
precipitation) after separation and concentration by HIC. HIC was
performed in a 15-mm 3 40-cm column packed with Phenol Sepharose
high-performance resin (Amersham Bioscience; Piscataway, NJ) equilibrated in 20 mM Mops, pH 7, 1 mM DTT, and 1 M ammonium sulfate. The
sample was adjusted to contain 20 mM Mops, pH 7, 1 mM DTT, and 1 M
ammonium sulfate and loaded directly onto the column. The column was
then washed in the same buffer and eluted with a 200-min, linear gradient
of 1.0 to 0 M ammonium sulfate with a flow rate of 4 mL/min. For initial
characterization of protease activity, all fractions were assayed with the
substrates Z-VAD-AFC and Z-DEVD-AFC. Two proteolytic activities were
identified: activity A eluting at 0.45 M ammonium sulfate and activity B
eluting at 0.3 M. For the purification protocol, the sample obtained from
the ammonium sulfate precipitation step was loaded and ran on the HIC
column as described above. Throughout the purification protocol,
fractions were assayed for Z-VAD-AFC hydrolytic activity.
Heparin Affinity Chromatography
Active fractions from the HIC column were pooled, and the buffer was
exchanged with 20 mM Mops, pH 7, 1 mM DTT, and 50 mM NaCl by
filtering through an Amicon Ultra-15, 30,000 MWCO centrifugal filter
device (Millipore, Bedford, MA). After buffer exchange, the sample was
loaded onto a 5-mL Heparin HP affinity column (Amersham Bioscience),
washed with 20 mM Mops, pH 7, 1 mM DTT, and 50 mM NaCl, and eluted
in 20 mM Mops, pH 7, 1 mM DTT, and 2 M NaCl.
Anion Exchange Chromatography
Active fractions from the heparin column were prepared for anion
exchange chromatography by replacing the buffer with 50 mM TrisHCl, pH 8.5, 2 mM EDTA, and 1 mM DTT (by concentration and washing in
an Amicon Ultra-15). The resulting sample was loaded onto a Mono Q
fast-protein liquid chromatography 5/5 column (Amersham Bioscience)
and eluted with a 200-min linear gradient of 0 to 0.3 M NaCl with a flow
rate of 0.5 mL/min. Two peaks of proteolytic activity eluted from the Mono
Q column and were combined into two separate fractions called SAS-1
and SAS-2 (see Results).
Size Exclusion Chromatography
The two active fractions from the Mono Q column were individually
concentrated with an Amicon Ultra-15 into 4 mL with the buffer consisting
of 20 mM Mops, pH 6.5, 1 mM DTT, and 150 mM NaCl. Each 4-mL sample
was loaded onto a Sephacryl S-200 (16/60) column (Amersham Bioscience) and run at 0.5 mL/min. Fractions containing proteolytic activity
(Z-VAD-AFC) were checked for purity by silver staining of an SDS-PAGE.
Substrate, Inhibitor, pH, and Salt Profiles
Samples of SAS-1 and SAS-2 from the Mono Q column were used for the
substrate, inhibitor, pH, and salt profile assays. Conditions were as
described above for substrate and inhibitor assays and as described
Caspase-Like Ser Proteases
below for the pH and salt assays. All activity assays were performed in
triplicate.
The optimal pH for hydrolytic activity against the substrate Z-VAD-AFC
was measured over the pH range of 5.5 to 7.5. A 20-mM Mes buffer
containing 0.5 M NaCl was used for the assays at pH 5.5, 6.0, and 6.5, and
20 mM Mops and 0.5 M NaCl were used for assays at pH 6.5, 7.0, and 7.5.
Five salts, NaCl, KCl, MgCl2, CaCl2, and ZnCl2, were tested at 10 mM
and 100 mM in 20 mM Mops, pH 6.5, or at 10, 100, and 500 mM in 20 mM
Mops, pH 6.5, for their affect on hydrolytic activity against Z-VAD-AFC.
871
ACKNOWLEDGMENTS
We thank Lilo Barofsky for her help with the mass spectrometry analysis
and Donald Buhler for the use of the fluorometer. We are also grateful to
Jennifer Lorang, Teresa Sweat, and Marc Curtis for their scientific
discussion and technical assistance. This work was supported in part by
grants from the USDA National Research Initiative Competitive Grants
Program (1999-02404 and 2001-35319-10896) and the National Science
Foundation (IBN-9631442).
Polyclonal Antibody Production
Received October 6, 2003; accepted January 27, 2004.
Polyclonal antibodies against SAS-1 and SAS-2 were produced at the
Laboratory of Animal Research, Oregon State University by subcutaneous injection in rabbits. After anion exchange chromatography, SAS-1
and SAS-2 (1 mg) were gel purified and mixed with the adjuvant
TiterMax Gold (Sigma) before injection. Booster injections (1 ug of
SAS-1 and SAS-2 mixed in TiterMax Gold) were administered at weeks
4 and 8, and final serums were collected at week 11.
Silver Staining and Protein Blotting
All enzyme samples were analyzed on an 8% SDS-PAGE. Silver staining
was performed following the protocol developed by Blum et al. (1987).
Electroblotting was done in a Trans-Blot SD semi-dry transfer cell (BioRad, Hercules, CA) with transfer buffer consisting of 25 mM Tris, 192 mM
glycine, and 0.02% SDS in 20% methanol. Proteins were blotted onto
a Protran nitrocellulose membrane (Keene, NH) with a 0.45-mm pore size
(Schleicher and Schuell).
Membranes were blocked with 1% BSA in TBST (100 mM Tris-HCl, pH
7.6, 0.9% NaCl, and 0.1% Tween 20) for 20 min. For protein gel blots,
SAS-2 polyclonal antibody was added at a 1:2000 dilution in TBST for 1 h
and washed three times for 15 min each in 13 TBST, and then 28 antibody
(goat anti-rabbit conjugated to horseradish peroxidase) (Santa Cruz
Biotechnology, Santa Cruz, CA) was added at a 1:100,000 dilution for 1 h.
Blots were washed four times for 15 min each in TBST before
development with SuperSignal West Pico chemiluminescent substrate
(Pierce, Rockford, IL). Biotin blots were incubated with NeutrAvidin
conjugated to horseradish peroxidase (Pierce) at a 1:50,000 dilution for
1 h, washed four times for 15 min each, and developed the same as
protein gel blots.
Amino Acid Sequencing and Mass Spectrometry
SAS-1 and SAS-2 samples from the Mono Q column were run in an SDSPAGE, blotted onto a polyvinylidene difluoride membrane, and stained
with 0.1% Coomassie Brilliant Blue R 250 in 1% acetic acid and 40%
methanol. The 84-kD bands were excised and destained in 50%
methanol. NH3-terminal sequencing was conducted by Edmonds
degradation at The Institute of Molecular Biology, University of Oregon
(Eugene, OR).
Purified SAS-1 and SAS-2 was analyzed by MALDI-TOF mass
spectrometry at the Mass Spectrometry Laboratory, Oregon State
University (Corvallis, OR).
The peptide maps of SAS-1 and SAS-2 were produced at the Stanford
PAN Biotechnology Facility, Stanford University (Stanford, CA). Samples
were excised from an SDS-PAGE, partially digested in-gel by trypsin,
separated by high-performance liquid chromatography, and analyzed by
MALDI-TOF mass spectrometry. Three internal peptides of SAS-2
produced by the in-gel tryptic digest were NH2-terminally sequenced
by Edmonds degradation at the Stanford PAN Biotechnology Facility.
Sequence data from this article have been deposited with the EMBL/
GenBank data libraries under accession numbers BAB89803, CAA76724,
CAA76725, T06577, NP_569048, and A55800.
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Purification and Characterization of Serine Proteases That Exhibit Caspase-Like Activity and Are
Associated with Programmed Cell Death in Avena sativa
Warren C. Coffeen and Thomas J. Wolpert
Plant Cell 2004;16;857-873; originally published online March 12, 2004;
DOI 10.1105/tpc.017947
This information is current as of June 18, 2017
References
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