©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
The Membrane of Leaf Peroxisomes Contains a Porin-like Channel (*)

(Received for publication, February 14, 1995; and in revised form, April 27, 1995)

Sigrun Reumann (1)(§), Elke Maier (2), Roland Benz (2), Hans W. Heldt (1)

From the  (1)Institut für Biochemie der Pflanze, Universität Göttingen, Untere Karspüle 2, D-37073 Göttingen and the (2)Lehrstuhl für Biotechnologie, Theodor-Boveri-Institut (Biozentrum) der Universität Würzburg, Am Hubland, D-97074 Würzburg, Federal Republic of Germany

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

Spinach leaf peroxisomes were purified by Percoll density gradient centrifugation. After several freeze-thaw cycles, the peroxisomal membranes were separated from the matrix enzymes by sucrose density gradient centrifugation. The purity of the peroxisomal membranes was checked by measuring the activities of marker enzymes and by using antibodies. Lipid bilayer membrane experiments with the purified peroxisomal membranes, solubilized with a detergent, demonstrated that the membranes contain a channel-forming component, which may represent the major permeability pathway of these membranes. Control experiments with membranes of other cell organelles showed that the peroxisomal channel was not caused by the contamination of the peroxisomes with mitochondria or chloroplasts.

The peroxisomal channel had a comparatively small single channel conductance of 350 pS in 1 M KCl as compared with channels from other cell organelles. The channel is slightly anion selective, which is in accordance with its physiological function. The single channel conductance was found to be only moderately dependent on the salt concentration in the aqueous phase. This may be explained by the presence of positive point net charges in or near the channel or by the presence of a saturable binding site inside the channel. The possible role of the channel in peroxisomal metabolism is discussed.


INTRODUCTION

Leaf peroxisomes belong to the microbodies, a group of small multipurpose cell organelles which are found in all, except some very primitive, eukaryotic cells(1) . They are surrounded by a single membrane. Peroxisomes usually, although not always, contain HO producing enzymes and catalase to eliminate the HO(2) . They all contain the enzymes for -oxidation of fatty acids. Present evidence suggests that peroxisomes are not formed de novo but grow and divide like plastids and mitochondria, although they have no genome of their own. There are indications that all peroxisomes, e.g. from fungi, higher plants, and animals, have a common origin, possibly an endosymbiotic event(2) .

Leaf peroxisomes play a vital role in photosynthesis. They are involved in the recycling of glycolate formed as an unavoidable by-product of CO fixation due to oxygen reacting instead of CO with ribulose bisphosphate. In a leaf the ratio of oxigenation/carboxylation during photosynthesis is between 0.2 and 0.5, resulting in very high metabolic fluxes through the leaf peroxisomes. The peroxisomal reaction chains are strictly compartmentalized. When leaf peroxisomes were subjected to an ``osmotic shock'' the peroxisomal membrane was damaged extensively, but surprisingly the peroxisomal matrix did not disintegrate and its metabolic function, including the high compartmentation of metabolism, was unaltered(3, 4) . The intermediates of the reaction chains did not leak out(5) . From these findings we concluded that the compartmentation of metabolism in leaf peroxisomes is not due to a boundary function of the peroxisomal membrane but is the result of the properties of the peroxisomal matrix which allow metabolite channelling(3) . Our results suggest that specific translocators are not essential for the functional compartmentation of metabolism. Nonspecific pores seem sufficient for the metabolite transfer into and out of the leaf peroxisomes. General diffusion channels, called porins, exist in the outer membrane of Gram-negative eubacteria (for review, see (6) ), mitochondria(7) , and plastids(8, 9) . Bacterial porins occur as trimers of three identical subunits. Each subunit contains one diffusion channel for small molecules. They are formed entirely of amphipathic -sheets arranged in a barrel-like structure(6, 7) .

The results are conflicting as to whether other types of peroxisomes contain a pore-forming protein or specific translocators. Liver peroxisomes were found to be permeable to sucrose and nucleotides(10) . Upon reconstitution experiments a channel forming activity was attributed to a 22-kDa membrane popypeptide(11) , but the subsequent analysis of the amino acid sequence did not show any similarity to known porin structures(12) . Other experiments incorporating membrane preparations of liver peroxisomes into liposomes, using the patch clamp technique(13) , and into planar lipid bilayers (14) indicated that large cation-selective voltage-gated pores with an estimated diameter of 1.5-3.0 nm may be responsible for the high permeability of liver peroxisomes. In the yeast Hansenula polymorpha, a 31-kDa peroxisomal integral membrane protein, with a structure similar to the 31-kDa mitochondrial porin of Saccharomyces cerevisiae, was claimed to be responsible for the in vitro permeability of the peroxisomes(15) .

Some results are not consistent with the presence of a free diffusion channel in peroxisomes. Yeast peroxisomal membranes contain a proton-translocating ATPase(16) , and a proton gradient across the peroxisomal membrane has been observed(17) . An ATPase activity was also found in the membranes of liver peroxisomes(18) . An integral membrane protein showing a high structural homology with mitochondrial anion translocators (19) has been identified in the peroxisomal membrane of the yeast Candida boidinii. This suggests that these peroxisomes contain a specific metabolite translocator. The boundary function of the peroxisomal membrane is thus still a matter of debate.

We set out to investigate whether leaf peroxisomal membranes contain channel forming activity. This report shows that leaf peroxisomal membranes contain channels allowing the passage of the metabolites of the photorespiratory metabolism and which are distinctly different from the porin channels of leaf mitochondria and chloroplasts.


EXPERIMENTAL PROCEDURES

Plant Material

Peroxisomes, mitochondria, and chloroplasts were isolated from leaves of spinach (Spinacia oleracea L., U.S Hybrid 424; Ferry-Morse Seed Company, Mountain View, CA). The plants were grown and harvested as described(4) .

Isolation of Leaf Peroxisomes

Peroxisomes were isolated by modifying to the method of Yu and Huang(20) . The scale was increased by a factor of five up to 350 g of leaves, and the homogenization procedure was intensified. The final peroxisomal pellet was resuspended and stored at -20 °C. The yield of peroxisomes was about 6 mg of protein with a specific activity of hydroxypyruvate reductase of about 19 µmol(minmg) (see Table 1). The intactness measured as latency of hydroxypyruvate reductase (3) was about 95%.



Isolation of Leaf Peroxisomal Membranes

The highly aggregated structure of the matrix enzymes was destroyed by subjecting the organelles (12 mg of protein) to five freeze-thaw cycles (freezing in liquid nitrogen, thawing at room temperature) and intensive homogenization in a Potter homogenizer. The suspension was adjusted to 30% (w/w) sucrose, loaded on a linear sucrose gradient (35-60% (w/w) sucrose in 10 mM HEPES, pH 7.5, 0.8 mM MgCl), and centrifuged in a swing-out rotor (240,000 g, 15 h, Sorvall AH 650). The gradient was fractionated from the top (12 fractions of 0.4 ml). The fractions were diluted to a sucrose content of 30% (w/w) and stored at -80 °C. All the data of enzyme activities (Fig. 1), SDS-PAGE()and Western blot analysis (Fig. 2), and porin activity (Fig. 6) were obtained with the same gradient and confirmed by additional experiments.


Figure 1: Subfractionation of the peroxisomal suspension on a sucrose gradient. The gradient fractions were analyzed for protein and sucrose concentration (A) and enzyme activity of the peroxisomal matrix enzymes hydroxypyruvate reductase (HPR) and catalase (CAT) (B), the peroxisomal membrane enzyme ACS (C), and cytochrome c reductase (CCR) and ferricyanide reductase (FR) (D). Results are expressed as percentage of total gradient activity in each fraction. Fractions 1 and 12 represent the top and the bottom fractions, respectively. Recoveries of protein and marker enzymes (recovery, total gradient protein, or activity) were checked (protein: 101%, 11.7 mg; hydroxypyruvate reductase: 80%, 114 µmolmin; catalase: 91%, 11.3 mmolmin; ACS: 84%, 128 nmolmin; NADH-ferricyanide reductase: 71%, 3.64 µmolmin; NADH-cytochrome c reductase: 79%, 44.2 nmolmin.




Figure 2: Western blot analysis as control for membrane contamination of the peroxisomal membrane fractions and SDS-PAGE. Fractions of the sucrose gradient were subjected to SDS-PAGE (A) and immunoblotted with a polyclonal antibody against the 24-kDa protein of the outer envelope membrane of spinach chloroplasts (B) or with a polyclonal antibody against the 30-kDa porin of pea root plastids, showing strong cross-reactivity with the 30-kDa porin of the outer membrane of spinach mitochondria (C) as described under ``Experimental Procedures.'' For Western blot analysis in each lane, 20 µg of protein and for silver stain 3 µg of protein were separated by SDS-PAGE. Fractions 8+9 and 10+11 were pooled in a protein ratio of 1:1 because of their minor protein content. As a control 5 and 20 µg of protein of spinach chloroplast envelope membranes and mitochondrial membranes were blotted. P, peroxisomal suspension.




Figure 6: Comparison of porin activity and content of peroxisomal membrane in the fractions of the sucrose gradient. The concentration of peroxisomal membrane was measured on the basis of the specific ACS activity. The channel-forming activity was determined as explained in the text by using membranes from diphytanoyl phosphatidylcholine/n-decane. The voltage applied was 10 mV; T = 25 °C.



Isolation of Chloroplast Envelopes and of Mitochondrial Membranes

Chloroplasts were isolated according to Heldt and Sauer (21) and chloroplast envelope membranes according to Douce et al.(22) .

Mitochondria were purified according to the method of Neuburger et al.(23) . The mitochondria were disrupted osmotically by incubation for 30 min in distilled water on ice and sedimented afterward (160,000 g, 45 min, Kontron TFT 65.13).

Measurement of Enzyme Activities

If not stated otherwise, measurements of marker enzyme activities were carried out at 25 °C in a final volume of 1 ml. Hydroxypyruvate reductase and catalase were measured as described previously(3) . Acyl-CoA synthetase (ACS) was measured as described by Fischer et al.(9) . The reaction was stopped after incubation for 10 min at 30 °C. The measurement of NADH- ferricyanide reductase (700 µl)(24) , NADH-cytochrome c reductase(25) , cytochrome c oxidase (700 µl) (24) , and NADP-glycerinaldehyde-3-phosphate dehydrogenase (26) was performed as described. Protein was determined according to Peterson (27) .

SDS-PAGE and Western Blot Analysis

Precipitation of protein fractions was carried out using chloroform-methanol(28) . SDS-PAGE was performed on 12.5% gels according to Laemmli(29) . Polypeptide bands were made visible by silver staining(30) . In Western blot analysis the bound antibodies were made visible with a peroxidase-coupled second antibody using an ECL-kit (Amersham-Buchler, Braunschweig, Germany).

Lipid Bilayer Experiments

The methods used for the bilayer experiments have been described in detail(31, 32) . Peroxisomal membranes were solubilized in 0.5% Genapol X-80 (Fluka, Neu-Ulm) and added to the aqueous phases at one or both sides of the black membranes.


RESULTS

Preparation of Peroxisomes

From measurement of the activities of hydroxypyruvate reductase and catalase as marker enzymes for the peroxisomal matrix, the yield of the peroxisomes obtained from spinach leaves was evaluated as about 5 and 11% (Table 1). This low yield reflects the difficulties of peroxisomal isolation and is inherent to all published preparation procedures(20, 33) . The contamination of the peroxisomal preparation with chloroplasts, mitochondria, and ER is quite low, as the recoveries of the corresponding marker enzyme activities were 0.05, 0.2, and 0.01% as compared to the starting homogenate (Table 1). From this the contamination of the peroxisomal fraction by mitochondria, chloroplasts, and ER can be evaluated as less than 1, 4, and 0.2%, respectively. The purity of the peroxisomal suspension has been checked earlier by electron microscopy(3) .

Isolation and Purification of the Peroxisomal Membrane

The most used method for the isolation of peroxisomal membranes of rat liver (11, 34) and glyoxysomes (35, 36) is treating the peroxisomes with 100 mM sodium carbonate at pH 11.5(37) . This method was unsuitable because our measurements indicated a destruction of the porin activity. After various attempts to solubilize the highly aggregated matrix enzymes, we found it best to disrupt the peroxisomes mechanically (see ``Experimental Procedures'') and to separate the membranes by centrifugation in a sucrose density gradient. At present, unfortunately, there is no specific marker enzyme for the peroxisomal membrane of leaves. ACS, which has been shown to be closely associated with the membranes of leaf peroxisomes (38) and glyoxysomes (a differentiation form of plant peroxisomes), is present in the outer membrane of the chloroplast envelope (39, 40, 41) and also in the microsomal membranes(42, 43, 44) . It is possible that the outer membranes of plant mitochondria also contain this enzyme(45) . The results of Table 1show that only a minor portion of the cellular ACS activity is associated with the peroxisomes. Despite this, because of the very low contamination of the peroxisomal suspension by other ACS-containing membranes (Table 1) the ACS could be used as marker for the peroxisomal membrane. If 50% of the ACS activity were associated with the chloroplasts, the ACS activity from contaminating chloroplasts in the peroxisomal preparation would be less than 25% of the measured ACS activity. The same calculation with ER results in 1%.

In the experiment shown in Fig. 1, the activities of marker enzymes for the various subcellular components have been measured in the different fractions of the sucrose density gradient. Most of the protein (about 80%) and the marker enzyme for the peroxisomal matrix catalase and hydroxypyruvate reductase are found in the upper part of the gradient. These fractions represent the peroxisomal matrix proteins. The ACS activity forms a distinct peak at a density of 1.21-1.23 g/ml. Control experiments by measuring specific marker enzymes and Western blot analysis of all fractions of the sucrose gradient excluded that this ACS peak resulted from nonperoxisomal sources. 1) One of the major constituents of the outer envelope membrane of spinach chloroplasts is a 24-kDa protein which function is unknown until now(46) . Using an antibody against this protein, we could localize the contaminating outer envelope membrane in the first fraction of the sucrose gradient (Fig. 2B) at its low equilibrium density concurring with earlier results(39, 47) . The 24-kDa protein was not detectable in the fractions containing ACS activity indicating that the outer envelope membrane is absent in these fractions. 2) NADH-cytochrome c reductase, a marker enzyme for ER membranes(25) , was found at low activities on the top of the gradient at a density of 1.15 g/ml as shown earlier(25) . 3) The outer membrane of mitochondria was detected with the marker enzyme NADH-ferricyanide reductase (22) and in addition by using polyclonal antibodies against the 30-kDa porin of non-green pea root plastids(9) . As the porins of plant non-green plastids and mitochondria are relatively homologous proteins(9) , these antibodies show cross-reaction with the mitochondrial 30-kDa porin of spinach (Fig. 2C) and are thus a useful marker for the detection of the outer membrane of spinach mitochondria. Using both methods we were able to show that the content of the outer mitochondrial membrane in that part of the gradient where the maximal ACS activity is localized is rather low. The cross-reaction with a polypeptide of 66 kDa in fractions 10-12 could be an nonspecific artifact. The activity of NADH-ferricyanide reductase in the upper part of the gradient (fractions 1-4) may be due to peroxisomal activity as potato tuber peroxisomes were shown to possess this enzyme(48) . Thylakoid membranes were enriched at 1.17-1.18 g/ml (data not shown) as reported earlier(49) .

As the ACS peak contained neither outer chloroplast envelope membranes, outer mitochondrial membranes, nor ER membranes to any detectable extent, it can be concluded that the ACS peak represents the peroxisomal membrane and that this membrane is not contaminated with outer membranes of chloroplasts or mitochondria to any appreciable amount. As the outer mitochondrial and outer chloroplast envelope membranes both contain porins, the absence of these membranes in the peroxisomal membrane fraction is essential. In gradients with incompletely disrupted peroxisomes, the peroxisomal membrane could be identified from the adhering activities of the peroxisomal enzymes catalase, hydroxypyruvate reductase, and malate dehydrogenase, forming a second smaller peak at 1.215 g/ml. Glyoxysomal membranes have been reported to equilibrate at this density(50) . SDS-PAGE was performed with the fractions 1-12 of the same gradient (Fig. 2A). The dominant proteins of the peroxisomal membrane had a subunit molecular mass of about 50, 48, 45, 43, 39, 32, and 13 kDa.

Solubilized Peroxisomes Show a Pore Forming Activity

In further experiments we investigated whether peroxisomes contain any pore forming activity. The sedimented peroxisomes (protein concentration about 1 mg/ml) were treated with the detergent Genapol X-80 (final concentration 0.5%) to solubilize the peroxisomal membrane. The detergent extract was added to the aqueous phase, bathing a lipid bilayer, and the membrane current was measured. Fig. 3demonstrates that there is indeed a pore forming activity in these detergent extracts. Each conductance step of Fig. 3corresponds to the incorporation of one channel-forming unit into the membrane. The average single channel conductance of these channels is only 350 pS in 1 M KCl (see the histogram in Fig. 4). The occurrence of a single channel conductance of 600 pS probably indicates the incorporation of a dimer. The conductance of the peroxisomal channel is rather small as compared with those channels formed by mitochondrial or chloroplast porins under otherwise identical conditions (see also below).


Figure 3: Single channel recording of a diphytanoyl phosphatidylcholine/n-decane membrane in the presence of detergent-solubilized spinach leaf peroxisomes. 20 min after the formation of the membrane 1 µg/ml detergent-solubilized spinach leaf peroxisomes was added to the aqueous phase on one side of the membrane. The aqueous phase contained 1 M KCl (pH 6). The applied membrane potential was 10 mV; T = 25 °C.




Figure 4: Histogram of the probability of the occurrence of certain conductivity units observed with membranes formed of diphytanoyl phosphatidylcholine/n-decane in the presence of 1 µg/ml detergent-solubilized spinach leaf peroxisomes. The aqueous phase contained 1 M KCl. The applied membrane potential was 10 mV; T = 25 °C. The average single channel conductance was 350 pS for 344 single channel events. The data were collected from 10 different membranes. P(G), probability of the single channel conductance G.



Identification of the Channel in the Peroxisomal Membrane

The channel formed by the detergent extracts from whole peroxisomes had a completely different single channel conductance as compared with the porins from other plant cell membranes. To give further direct evidence that the origin of the novel channel-forming protein is the peroxisomal membrane, we investigated the channel forming activity of the various fractions of the sucrose gradient as follows: small quantities of the fractions were dissolved in Genapol X-80 and added to the aqueous phase on both sides of an artificial bilayer (final protein concentration in the aqueous phase 0.6 µg/ml). After a lag time of a few minutes, probably caused by slow aqueous diffusion of the protein, the conductance of the membrane, caused by the insertion of channels into the membrane, started to increase. The time course of the increase was similar to that described previously for porins of mitochondrial or bacterial origin(51, 52) . The number of inserted pores with a single channel conductance of less than 1 nS in a given time (20 min) was counted. The distribution of these conductance steps in a histogram (data not shown) was similar to that shown for whole peroxisomes (Fig. 4). The mean values of three measurements were taken as a semiquantitative measure for the peroxisomal channel forming activity of the sample. After the insertion of more than 30-50 pores in one black membrane, the single channel conductance could not be clearly identified (see Fig. 3and Fig. 5). As the channel incorporation was so frequent for the fractions containing the peroxisomal membranes, several bilayers had to be painted during the measuring period. Control experiments showed that the addition of the detergent Genapol X-80 alone at a similar concentration to that used with the protein did not lead to any appreciable increase in the membrane conductance.


Figure 5: Channel formation by peroxisomal membranes taken from a sucrose density gradient. Stepwise increase of the membrane current given after the addition of Genapol-solubilized peroxisomal membranes (A) to a black lipid bilayer membrane given as a function of time and comparison with chloroplast envelope (B) and mitochondrial porin activity (C). In A the peroxisomal membrane fraction of the sucrose gradient was added. Sometimes the incorporation of larger pores, similar to described chloroplast envelope porins (B) and mitochondrial porins (C), was observed. The aqueous phase contained about 1 µg/ml protein and 1 M KCl. The membrane was formed from diphytanoyl phosphatidylcholin/n-decane. The voltage applied was 10 mV; T = 25 °C.



We were able to identify three different types of channels in the protein samples taken from the sucrose density gradient. The light fractions contained (besides the 350 pS channel; see Fig. 5A) a channel with a giant single channel conductance of 7-9 nS in 1 M KCl. This channel was similar to that found in reconstitution experiments with the outer membrane of the chloroplast envelope (see Fig. 5C; 8). In other fractions (preferentially fractions 2-3), we sometimes observed an additional channel, which was indistinguishable from mitochondrial porin from plant and other sources (see Fig. 5B; 7). The most prominent channel, however, in all fractions was the 350 pS channel. Fig. 6shows the distribution of this channel within the fraction of the sucrose density gradient. It is noteworthy that the peroxisomal porin activity measured as the number of channels (20 min after the protein addition) in a membrane with a surface area of 1 mm corresponded to the 350 pS channel only. When one of the other channels did happen to incorporate into the lipid bilayer membrane, the experiment was stopped and a new experiment was started. This was necessary because of the much higher single channel conductance of the mitochondrial and chloroplast porins. Fig. 6shows also the specific activity of ACS, i.e. the content of peroxisomal membrane in the fractions of the sucrose density gradient. There is a correlation between the specific ACS activity and the pore forming activity showing that the 350 pS channel resides within the peroxisomal membrane.

Properties of the Peroxisomal Channel

After the addition of whole peroxisomes or peroxisomal membranes, which had been solubilized with Genapol X-80, step increases in membrane conductance could be resolved (see Fig. 3and Fig. 5). The average single channel conductance with 1 M KCl was 350 pS (see the histogram of Fig. 4). Fig. 3and Fig. 5show also that most of the steps were directed upwards. Only a few downward steps were observed which means that the peroxisomal channels had a long lifetime. Even at higher transmembrane potentials of about 50 mV the closing events did not become more frequent. This demonstrates that the peroxisomal porin is not voltage-regulated at these potentials.

The channel formed by peroxisomal porin was permeable for a variety of ions. Table 2summarizes the single channel conductance for different salt solutions. The nature of the anions had a substantial influence on the single channel conductance, whereas the influence of the cations was rather small. Thus, the single channel conductance in 1 M KCl was approximately the same as in 1 M LiCl, but it was considerably smaller in 1 M potassium acetate (K and Cl and Li and acetate have the same aqueous mobility)(53) . This result suggests that the channel had a certain preference for anions. We found that a variety of organic anions, such as formiate, glycerate, and glycolate are permeating this channel. It is noteworthy, that the conductance of the channel was not a linear function of the bulk aqueous concentration, which may indicate that positive point net charges are localized in or near the channel mouth. On the other hand, it is also possible that the channel contains a binding site for organic anions, which facilitates the diffusion of these ions in a similar way as specific bacterial porins do for certain substrates such as sugars, nucleosides, and phosphate(6) .



Ion Selectivity

The single channel data suggested that the peroxisomal channel is anion-selective. We performed zero current membrane potential measurements to study its ion selectivity in more detail. Membranes were formed in 100 mM KCl, and detergent-solubilized peroxisomal membrane was added to the aqueous phase when the membranes were in the black state. After incorporation of 100-1000 channels into a membrane, salt gradients were established by addition of small amounts of 3 M KCl solution to one side of the membrane. Under these conditions, the more diluted side of the membrane became negative which indicated indeed preferential movement of chloride over potassium. However, the potential and the permeability ratio between potassium and chloride (as calculated from the Goldman-Hodgkin-Katz equation; 54) differed considerably from experiment to experiment, which probably means that other channels such as mitochondrial porin or the chloroplast porin interfered with the measurements and led to irreproducible results. This is because the single channel conductance of the peroxisomal porin (350 pS) is about 6-12 times smaller than that of mitochondrial porin (2.4 or 4.0 nS) and about 23 times smaller than that of chloroplast porin (8 nS) under otherwise identical conditions (1 M KCl).


DISCUSSION

The Peroxisomal Channel Has Properties Different from Channels of Other Cell Organelles

The peroxisomal membrane of leaf peroxisomes contains a channel-forming protein that is different from those of other cell organelles. The channel shows a single channel conductance of 350 pS (1 M KCl) which is much lower than those formed by the porins of mitochondria and chloroplasts under otherwise identical conditions (see Table 3). Furthermore, the single channel conductance in salts containing organic anions is much smaller as expected from the mobility of these ions in the aqueous phase. Apparently the peroxisomal channel is not a wide, water-filled channel like the channels of the mitochondrial and the chloroplast porins. The single channel conductance is not a linear function of the bulk aqueous concentration, caused either by positive net charges in or near the channel mouth or by a binding site for anions. This is another indication that the peroxisomal channel has specialized channel properties. The size of the peroxisomal channel is somewhat difficult to obtain from the single channel measurements since only the size of wide, water-filled channels may be obtained from their conductance(6, 7) . As shown in Table 2the K salts of formiate, acetate, glycerate, and glycolate are able to penetrate the channels, but the channel conductance with these ions is much smaller than with KCl. It appears from this result that the channels are just large enough to let these organic anions pass through. From the size of the permeating molecules, a channel diameter of about 1 nm may be estimated. The channel appears to be well suited to enable the transfer of metabolites in and out of the peroxisomes during photorespiratory metabolism.



Do Other Peroxisomal Membranes Also Contain an Ion-permeable Channel?

Our data strongly suggest that plant peroxisomes contain an ion-permeable channel. The question arises whether pores are a common principle of the permeability properties of peroxisomal membranes. We mentioned in the Introduction that peroxisomes from yeast and liver may contain pores that have very similar properties to those in the outer membrane of the mitochondria(11, 12, 15) . In particular, the channels reconstituted from liver peroxisomes show the same high voltage dependence, commencing at about 20-30 mV, as mitochondrial porins(7, 11) . We did not observe any voltage dependence up to 70 mV, indicating that the channel from leaf peroxisomes described here is not voltage regulated within the physiological range of membrane potentials. Obviously the channels attributed to liver peroxisomes have completely different properties than the channel described in our study. It remains to be studied whether the relatively small peroxisomal pores characterized here to be completely different from mitochondrial and chloroplast pores are a special feature of leaf peroxisomes, or whether they are also present in peroxisomes from other cells.


FOOTNOTES

*
This work was supported by the Deutsche Forschungsgemeinschaft (to H. W. H.), Project B9 of the Sonderforschungsbereich 176, and by the Fonds der Chemischen Industrie. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence should be addressed: Institut für Biochemie der Pflanze, Universität Göttingen, Untere Karspüle 2. D-37073 Göttingen, Germany. Tel.: +49-0551-399368; Fax: +49-0551-395749.

The abbreviation used is: PAGE, polyacrylamide gel electrophoresis.


ACKNOWLEDGEMENTS

We thank Dr. S. Borchert for useful discussions and M. Raabe for her excellent help in preparation of peroxisomes. The antibodies were kindly provided by Prof. Dr. Flügge (Universität Köln).


REFERENCES
  1. Cavalier-Smith, T. (1987)Nature 326, 332-333 [CrossRef][Medline] [Order article via Infotrieve]
  2. Borst, P.(1989) Biochim. Biophys. Acta 1008, 1-13 [Medline] [Order article via Infotrieve]
  3. Heupel, R., Markgraf, T., Robinson, D. G., and Heldt, H. W.(1991)Plant. Physiol. 96, 971-979
  4. Reumann, S., Heupel, R., and Heldt, H. W.(1994)Planta 193, 167-173
  5. Heupel, R., and Heldt, H. W.(1994)Eur. J. Biochem. 220, 165-172 [Abstract]
  6. Benz, R. (1994) in Bacterial Cell Wall (Ghuysen, J.-R., and Hakenbeck, R., eds) pp. 397-423, Elsevier Science B.V., Amsterdam
  7. Benz, R.(1994) Biochim. Biophys. Acta 1197, 167-196 [Medline] [Order article via Infotrieve]
  8. Flügge, U. I., and Benz, R.(1984)FEBS Lett. 169, 85-89 [CrossRef]
  9. Fischer, K., Weber, A., Brink, S., Arbinger, B., Schünemann, D., Borchert, S., Heldt, H. W., Popp, B., Benz, R., Link, T. A., Eckerskorn, C., and Flügge, U. I.(1994)J. Biol. Chem. 269, 25754-25760 [Abstract/Free Full Text]
  10. van Veldhoven, P. P., Debeer, L. J., and Mannaerts, G. P.(1983)Biochem. J. 210, 685-693 [CrossRef][Medline] [Order article via Infotrieve]
  11. van Veldhoven, P., Just, W. W., and Mannaerts, G. P.(1987)J. Biol. Chem. 262, 4310-4318 [Abstract/Free Full Text]
  12. Kaldi, K., Diestelkötter, P., Stenbeck, G., Auerbach, S., Jäkle, U., Mägert, H. J., Wieland, F. T., and Just, W. W.(1993) FEBS Lett. 315, 217-222 [CrossRef][Medline] [Order article via Infotrieve]
  13. Lemmens, M., Verheyden, K., van Veldhoven, P., Vereecke, J., Mannaerts, G. P., and Carmeliet, E.(1989)Biochim. Biophys. Acta 984, 351-359 [Medline] [Order article via Infotrieve]
  14. Labarca, P., Wolff, D., Soto, U., Necochea, C., and Leighton, F.(1986)J. Membr. Biol. 94, 285-291 [Medline] [Order article via Infotrieve]
  15. Sulter, G. J., Verheyden, K., Mannaerts, G., Harder, W., and Veenhuis M.(1993) Yeast 9, 733-742 [Medline] [Order article via Infotrieve]
  16. Douma, A. C., Veenhuis, M., Sulter, G. J., and Harder, W.(1987)Arch. Microbiol. 147, 42-47 [Medline] [Order article via Infotrieve]
  17. Nicolay, K., Veenhuis, M., Douma, A. C., and Harder, W.(1987)Arch. Microbiol. 147, 37-41 [Medline] [Order article via Infotrieve]
  18. Kamijo, K., Taketani, S., Yokota, S., Osumi, T., and Hashimoto, T.(1990)J. Biol. Chem. 265, 4534-4540 [Abstract/Free Full Text]
  19. Jank, B., Habermann, B., and Schweyen, R. J.(1993)Trends Biochem. Sci. 18, 427-428 [Medline] [Order article via Infotrieve]
  20. Yu, C., and Huang, A. H. C.(1986)Arch. Biochem. Biophys. 245, 125-133 [Medline] [Order article via Infotrieve]
  21. Heldt, H. W., and Sauer, F.(1971)Biochim. Biophys. Acta 234, 83-91 [Medline] [Order article via Infotrieve]
  22. Douce, R., Holtz, R. B., and Benson, A. A.(1973)J. Biol. Chem. 248, 7215-7222 [Abstract/Free Full Text]
  23. Neuburger, M., Journet, E.-P., Bligny, R., Carde, J. P., and Douce, R.(1982) Arch. Biochem. Biophys. 217, 312-323 [Medline] [Order article via Infotrieve]
  24. Douce. R., Mannella, C. A., and Bonner, W. D., Jr.(1973)Biochim. Biophys. Acta 292, 105-116 [Medline] [Order article via Infotrieve]
  25. Sauer, A., and Robinson, D. G.(1985)Planta 166, 227-233
  26. Gerhardt, R., and Heldt, H. W.(1984)Plant Physiol. 75, 542-547
  27. Peterson, G. L. (1977)Anal. Biochem. 83, 346-356 [Medline] [Order article via Infotrieve]
  28. Wessel, D., and Flügge, U.-I.(1984)Anal. Biochem. 138, 141-143 [Medline] [Order article via Infotrieve]
  29. Laemmli, U. K. (1970)Nature 227, 680-685 [Medline] [Order article via Infotrieve]
  30. Blum, H., Beier, H., and Gross, H. J.(1987)Electrophoresis 8, 93-99
  31. Benz, R., Janko, K., Boos, W., and Läuger, P.(1978)Biochim. Biophys. Acta 511, 305-319 [Medline] [Order article via Infotrieve]
  32. Schmid, A., Krömer, S., Heldt, H. W., and Benz, R.(1992)Biochim. Biophys. Acta 1112, 174-180 [Medline] [Order article via Infotrieve]
  33. Schmitt, M. R., and Edwards, G. E.(1983)Plant Physiol. 72, 728-734
  34. Fujiki, Y., Fowler, S., Shio, H., Hubbard, A. L., and Lazarow, P. B.(1982)J. Cell Biol. 93, 103-110 [Abstract]
  35. Chapman, K. D., and Trelease, R. N.(1992)Plant Physiol. Biochem. 30, 1-10
  36. Wolins, N. E., and Donaldson, R. P.(1994)J. Biol. Chem. 269, 1149-1153 [Abstract/Free Full Text]
  37. Fujiki, Y., Hubbard, A. L., Fowler, S., and Lazarow, P. B.(1982)J. Cell Biol. 93, 97-102 [Abstract]
  38. Gerhardt, B. (1986)Physiol. Vég. 24, 397-410
  39. Andrews J., and Keegstra, K.(1983)Plant Physiol. 72, 735-740
  40. Block, M. A., Dorne, A. J., Joyard, J., and Douce, R.(1983)FEBS Lett. 153, 377-381 [CrossRef]
  41. Joyard, J., and Stumpf, P. K.(1981)Plant Physiol. 67, 250-256
  42. Lessire, R., and Cassagne, C.(1979)Plant Sci. Lett. 16, 31-39
  43. Sukumar, V., and Sastry, P. S.(1987)Biochem. Int. 14, 719-726
  44. Ichihara, K., Nakagawa, M., and Tanaka, K.(1993)Plant Cell Physiol. 34, 557-566
  45. Frentzen, M., Neuburger, M., Joyard, J., and Douce, R.(1990)Eur. J. Biochem. 187, 395-402 [Abstract]
  46. Fischer, K., Weber, A., Arbinger, B., Brink, S., Eckerskorn, C., and Flügge, U. I. (1994)Plant Mol. Biol. 25, 167-177 [Medline] [Order article via Infotrieve]
  47. Block, M. A., Dorne, A. J., Joyard, J, and Douce, R.(1983)J. Biol. Chem. 258, 13273-13280 [Abstract/Free Full Text]
  48. Struglics, A., Fredlund, K. M., Rasmusson, A. G., and M, I. M.(1993) Physiologia Plantarum 88, 19-28 [CrossRef]
  49. Ludwig, B., and Kindl, H.(1976)Hoppe-Seyler's Z. Physiol. Chem. 357, 177-186
  50. Preisig-Müller, R., Muster, G., and Kindl, H.(1994)Eur. J. Biochem. 219, 57-63 [Abstract]
  51. Roos, N., Benz, R., and Brdiczka, D.(1982)Biochim. Biophys. Acta 686, 204-214 [Medline] [Order article via Infotrieve]
  52. Benz, R., Ishii, J., and Nakae, T.(1980)J. Membr. Biol. 56, 19-29 [Medline] [Order article via Infotrieve]
  53. Castellan, G. W. (1983) Physical Chemistry, pp. 769-780, Addison-Wesley, Reading MA
  54. Benz, R., Janko, K., and Läuger P.(1979)Biochim. Biophys. Acta 551, 238-247 [Medline] [Order article via Infotrieve]
  55. Roos, N., Benz, R., and Brdiczka, D.(1982)Biochim. Biophys. Acta 686, 204-214 [Medline] [Order article via Infotrieve]
  56. Ludwig, O., Krause, J., Hay, R., and Benz, R.(1988)Eur. Biophys. J. 15, 269-276 [Medline] [Order article via Infotrieve]
  57. Aljamal, J. A., Genchi, G., De Pinto, V., Stefanazzi, L., De Santis, A., Benz, R., and Palmieri, F.(1993)Plant Physiol. 102, 615-621 [Abstract/Free Full Text]

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