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Quantitation showed that nearly five times the amount of protein was recovered in the first elution using the spin columns (Fig. 2B). Overall, the total extraction of protein combined from both elutions was about three times greater through the use of the spin columns. This result has been reproduced in our laboratory for a variety of GST-fusion proteins and their binding proteins, as well as for IPs (data not shown). To assess the reproducibility of protein recovery using pelleting or spin columns, a multireplicate experiment was performed. Poor reproducibility was commonly achieved using the pelleting method, despite two pooled elutions with 1⫻ SDS sample buffer (Fig. 2C, upper panel). In contrast, the spin column approach was highly reproducible, using only a single elution of protein in 1⫻ SDS sample buffer (Fig. 2C, lower panel). Densitometry was performed, and the results were normalized against the mean of the data set (given a value of 1). The standard deviation of the pelleting data was 0.22, compared with 0.05 for the spin column method. Thus, the spin column method was approximately four times more reproducible. The use of spin columns to perform pull-down assays and IPs presents a number of key advantages for analysis of protein–protein interactions by SDS–PAGE. (i) Threefold improvements in total protein recovery from beads. The increase probably arises from removal of a greater proportion of the volume of buffer within and around the beads that is normally inaccessible to a pipet tip or needle using the pelleting method. (ii) Greatly enhanced reproducibility of extraction. The reproducibility of protein extraction throughout a set of replicate samples also dramatically improves through the use of spin columns. This is most likely due to the retention of the beads within the spin column, eliminating the possibility that some beads (and thus protein) may be accidentally and inconsistently removed with the supernatant during the washing steps in the pelleting method. (iii) Significantly smaller sample volumes. Through the use of the spin column, almost all of the protein eluted from the beads is collected in the first elution, whereas the pelleting method had a significant amount of protein remaining in the elution buffer retained between the beads. This is of significance when the whole sample is to be applied to SDS– PAGE due to restrictions on volumes that can be loaded in sample wells. Therefore, the new method eliminates further time-consuming protein concentration steps (4). (iv) Faster sample processing times can be achieved with spin columns due to the shorter centrifugation times and the fact that the tubes are not individually handled for supernatant removal with a pipet tip or needle. In summary, we have developed a spin-column-based method for performing protein pull-down or IP experiments that is faster, improves protein recovery, decreases

sample volume, and increases reproducibility across a series of replicate samples for SDS–PAGE analysis. Acknowledgments. This work was supported by grants from the Australian National Health and Medical Research Council (NHMRC to P.J.R. and to B.D.R.). M.A.C. was supported by a long-term fellowship from the Human Frontiers of Science Program. Pietro DeCamilli is thanked for providing the amphiphysin II SH3-domain GST-fusion protein bacterial expression plasmid and Peter Rowe for critical reading of the manuscript.

REFERENCES 1. Coligan, J. E., Dunn, B. M., Ploegh, H. L., Speicher, D. W., and Wingfield, P. T. (Eds.) (1999) Current Protocols in Protein Science, pp. 19.4.1–19.4.9, Wiley, New York. 2. Wang, X., Bruderer, S., Rafi, Z., Xue, J., Milburn, P. J., Kramer, A., and Robinson, P. J. (1999) EMBO J. 18, 4549 – 4559. 3. Butler, M. H., David, C., Ochoa, G. C., Freyberg, Z., Daniell, L., Grabs, D., Cremona, O., and De Camilli, P. (1997) J. Cell Biol. 137, 1355–1367. 4. Wessel, D., and Flugge, U. I. (1984) Anal. Biochem. 138, 141–143.

An Assay for Phosphoinositide Phosphatases Utilizing Fluorescent Substrates Gregory S. Taylor and Jack E. Dixon 1 Department of Biological Chemistry, University of Michigan Medical School, Ann Arbor, Michigan 48109-0606 Received April 17, 2001; published online June 27, 2001

Phosphoinositides are key signaling molecules that serve as second messengers in many important cellular processes including growth, metabolism, membrane trafficking, motility, differentiation, adhesion, and programed cell death. Considerable attention has been focused on identifying the enzymes that regulate the synthesis, breakdown, and interconversion of inositol lipids to better understand their role in these processes. We have recently described a nonradioactive procedure for assaying phosphoinositide phosphatase activity that employs malachite green-based detection of inorganic phosphate (1). Although this procedure is both sensitive and quantitative, it is not always useful for determining enzyme specificity because it based on the measurement of inorganic phosphate rather than the phosphoinositide moiety. Herein we describe a simple, nonradioactive method for assaying phosphoinositide phosphatase activity utilizing fluorescent phosphoinositide substrates and 1 To whom correspondence should be addressed. Fax: (734) 7636492. E-mail: [email protected].

Analytical Biochemistry 295, 122–126 (2001) doi:10.1006/abio.2001.5179 0003-2697/01 $35.00 Copyright © 2001 by Academic Press All rights of reproduction in any form reserved.

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thin-layer chromatography (TLC).2 This approach offers several advantages over existing methodologies. By analyzing the phosphoinositide reaction product(s) rather than release of inorganic phosphate, phosphatase specificity toward distinct sites on the inositol ring can be assessed more effectively. In addition, the fluorescent substrates are water soluble, thus eliminating the need to remove organic solvents from lipid substrate stock solutions or sonicate lipids into aqueous suspensions. This method can be used with purified recombinant phosphatases or immunoprecipitates and provides an alternative for determining the enzyme specificity that is complementary to the sensitivity of the malachite greenbased system. Materials and Methods Expression and purification of recombinant myotubularin and PTEN. Recombinant myotubularin (MTM1) was expressed with a C-terminal six-histidine tag in Escherichia coli BL21(DE3) Codon Plus cells (Stratagene, La Jolla, CA). A cDNA fragment containing the MTM1 open-reading frame without a stop codon was amplified by PCR using oligonucleotide primers containing 5⬘-NheI and 3⬘-SalI restriction linkers. This fragment was inserted into pET21a vector digested with NheI and XhoI to create pET-MTM1H 6. His-tagged MTM1 was expressed and purified as described for the Sac1p fusion protein (1). Bacterial recombinant PTEN was expressed and purified as previously described (2). Immunoprecipitation of MTM1 and PTEN proteins. Wild-type and catalytically inactive (active site Cys to Ser) mutant MTM1 and PTEN proteins with N-terminal FLAG epitope tags were overexpressed in Cos 1 cells. Wild-type and C375S MTM1 and PTEN wild-type and C124S FLAG expression vectors were constructed as described previously (2, 3). Cos 1 cells were maintained at 37°C and 5% CO 2 in Dulbecco’s modified Eagle medium containing 10% fetal calf serum, 50 U/ml (each) penicillin and streptomycin, and 4 mM glutamine. Cos 1 cells were transiently transfected in 100-mm dishes (2 ⫻ 10 6 cells per dish) with 10 ␮g MTM1 or PTEN expression vector DNA using Fugene 6 transfection reagent (Roche Diagnostics, Indianapolis, IN) according to the manufacturer’s protocol. At 30 h posttransfection, cells were removed from the plates, washed once with phosphate-buffered saline, and lysed on ice in a buffer consisting of Tris-buffered saline containing 1% Triton X-100, 0.025% (v:v) 2-mercaptoethanol, 1 mM phenylmethylsufonyl fluoride, 1 mM benzamidine, 1 mM EDTA, and 1 ␮g/ml (each) aprotinin, leupeptin, and pepstatin. Lysates were 2 Abbreviations used: TLC, thin-layer chromatography; MTM1, myotubularin.

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cleared by centrifugation (20 min at 18,000g, 4°C) and the supernatants incubated on a rocker overnight at 4°C with 50 ␮l Anti-FLAG M2-agarose affinity resin (Sigma, St. Louis, MO). MTM1 and PTEN affinity resin samples were washed with 5 ⫻ 1 ml of lysis buffer (above), followed by 2 ⫻ 1 ml washes with their respective phosphatase reaction buffers containing 0.025% 2-mercaptoethanol rather than dithiothreitol (below). Phosphatase assays. All fluorescent phosphoinositide substrates used in this study were obtained from Echelon Research Laboratories (Salt Lake City, UT). Phosphatase assays using bacterial recombinant MTM1 were conducted in 20 ␮l of MTM1 reaction buffer containing 50 mM ammonium acetate (pH 6.0) and 2 mM dithiothreitol with 1.0 ␮g fluorescent di-C 6NBD6 phosphoinositide substrate for 20 min at 30°C. Phosphatase assays with bacterial recombinant PTEN were conducted (in duplicate) in 20 ␮l of PTEN reaction buffer containing 50 mM ammonium carbonate (pH 8.0) and 2 mM dithiothreitol with 1.5 ␮g di-C 6-NBD6phosphatidylinositol 3,4,5-trisphosphate for various times at 37°C. MTM1 and one set of PTEN phosphatase reactions were terminated by the addition of 100 ␮l of acetone and then evaporated to dryness in a Speed-Vac evaporator set on low heat. The duplicate set of PTEN reactions was terminated by the addition of 80 ␮l malachite green reagent and inorganic phosphate release quantitated as described (1). Phosphatase assays using FLAG-tagged MTM1 wild-type or C375S proteins bound to agarose affinity resin (⬃25 ␮l resin) were incubated for 30 min at 30°C in 30 ␮l of MTM1 reaction buffer (above) containing 1.5 ␮g di-C 6-NBD6-phosphatidylinositol 3-phosphate. At the indicated time, the samples were centrifuged 10 s to pellet the beads and 20 ␮l of supernatant was removed. Acetone (100 ␮l) was added to each supernatant, and the samples were dried as described above. Phosphatase assays using FLAG-tagged PTEN wildtype or C124S proteins bound to agarose affinity resin (⬃25 ␮l resin) were incubated for 30 min at 37°C in 30 ␮l of PTEN reaction buffer containing 1.5 ␮g di-C 16NBD6-phosphatidylinositol 3,4,5-trisphosphate. The reactions were terminated and processed as described for FLAG-tagged MTM1 proteins above. Thin-layer chromatography. The dried phosphatase assay reaction products were resuspended in 10 ␮l of methanol/2-propanol/glacial acetic acid (5/5/2) and spotted onto a glass-backed TLC plate (K6 silica gel 60 Å, 20 ⫻ 20 cm, Whatman Inc., Clifton, NJ). The TLC plate was developed in a solvent system consisting of chloroform/methanol/acetone/glacial acetic acid/water (70/50/ 20/20/20) as described previously and air-dried (4). Fluorescent lipids were visualized and quantitated by UV light using a NucleoVision Imaging Workstation with GelExpert software (NucleoTech Corp., San Carlos, CA).

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FIG. 1. Determination of myotubularin substrate specificity using fluorescent phosphoinositides. Phosphatase assays containing 1.0 ␮g of di-C 6-NBD6-phosphoinositide substrate (indicated below) were carried out for 20 min at 30°C with buffer (⫺) or 5 ng bacterial recombinant MTM1 (⫹). Reactions were terminated by the addition of 100 ␮l acetone and dried under vacuum. Samples were separated by TLC as described under Materials and Methods and the fluorescent phosphoinositide bands visualized under UV light. The migration positions of authentic di-C 6-NBD6-phosphoinositide standards are indicated (at left) and the origin and solvent front are shown (at right). The di-C 6-NBD6-phosphatidylinositol reaction product generated by MTM1 is indicated by a (white arrow).

Results and Discussion We and others have previously shown that myotubularin, a phosphoinositide phosphatase mutated in the human genetic disorder myotubular myopathy, is highly specific for phosphatidylinositol 3-phosphate (3, 5). To determine the usefulness of fluorescent phosphoinositides for assessing inositol lipid phosphatase specificity, recombinant myotubularin was tested using synthetic di-C 6-NBD6-fluorescent phosphoinositide substrates. Initial experimentation revealed that buffer salts normally present in phosphatase assays interfered with subsequent analysis by TLC. Thus, the volatile buffering agents ammonium acetate (MTM1 assays) or ammonium carbonate (PTEN assays), which were removed during the drying process, were subsequently used. Of the panel of substrates tested, only NBD6-PI(3)P was dephosphorylated by myotubularin, which converted it to NBD6-phosphatidylinositol as shown in Fig. 1 (white arrow). Myotubularin did not dephosphorylate other mono-, bis-, or tris-phosphorylated fluorescent phosphoinositide species as is evident in Fig. 1. This result demonstrates that fluorescent

phosphoinositide substrates can be effectively utilized to probe inositol lipid phosphatase specificity. Next, a time course of di-C 6-NBD6-phosphatidylinositol 3,4,5-trisphosphate dephosphorylation by the human tumor suppressor PTEN phosphoinositide phosphatase was performed in duplicate and the reaction products were quantitated by two different methods. One set of samples was analyzed utilizing a malachite green-based inorganic phosphate detection system (1). The second set of samples was processed by TLC as described under Materials and Methods. The amount of the di-C 6-NBD6-phosphatidylinositol 4,5-bisphosphate reaction product in each lane was determined from the scanned image by comparison to standards (Fig. 2A). A plot of the quantity of reaction product produced versus time is shown in Fig. 2B. Although the shapes of the two curves are similar, reaction progress as monitored by di-C 6-NBD6-phosphatidylinositol 4,5bisphosphate release was approximately 35 to 40% greater than that determined by release of inorganic phosphate at any time point. This discrepancy is most likely due to the inherent limitations in quantitation of

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expressed in Cos 1 cells as fusion proteins with N-terminal FLAG epitope tags and immunoprecipitated using anti-FLAG affinity resin. The FLAGtagged fusion proteins bound to the affinity resin were then tested for phosphatase activity toward fluorescent phosphoinositide substrates. As shown in Fig. 3, both myotubularin (Fig. 3A, right lane) and PTEN (Fig. 3B, right lane) overexpressed and immunoprecipitated from Cos 1 cells were active toward di-C 6-NBD6-phosphatidylinositol 3-phosphate and di-C 16-NBD6-phosphatidylinositol 3,4,5-trisphosphate, respectively. Control immunoprecipitates from cells expressing catalytically inactive myotubularin C375S (Fig. 3A, left lane) or PTEN C124S (Fig. 3B, left lane) mutant proteins exhibited no activity. Furthermore, levels of both wild-type and mutant myotubularin or PTEN proteins were indistinguishable by immunoblotting (not

FIG. 2. Time course of di-C 6-NBD6-phosphatidylinositol 3,4,5trisphosphate dephosphorylation by PTEN. Duplicate phosphatase assays with 1.5 ␮g di-C 6-NBD6-phosphatidylinositol 3,4,5-trisphosphate were carried out for the indicated time at 37°C with 2.1 ␮g bacterial recombinant PTEN. (A) One set of reactions was terminated by the addition of 100 ␮l acetone, dried under vacuum, and analyzed by TLC. Phosphoinositide bands visualized under UV light were quantitated using GelExpert software (Nucleotech Corp.). The migration position of authentic di-C 6-NBD6-phosphoinositide standards and the origin are shown (at left). (B) The second set of reactions was terminated by the addition of 80 ␮l malachite green reagent and inorganic phosphate release quantitated as previously described (1). The values obtained for release of di-C 6-NBD6-phosphatidylinositol 4,5-bisphosphate (open circles) and inorganic phosphate (closed circles) are shown (in pmol) as a function of time.

scanned images. Because the accuracy of inorganic phosphate determination using malachite green-based assay systems has been previously demonstrated by numerous researchers, these data indicate that only a rough approximation of lipid phosphatase activity can be made using fluorescent phosphoinositides with the procedure described here. A common difficulty in testing putative lipid phosphatases for activity is that bacterial expression and purification can be problematic, particularly with large proteins. In addition, posttranslational modifications that may affect phosphatase activity often do not occur in bacteria. Therefore, we wanted to assess whether proteins overexpressed and immunoprecipitated from mammalian cells could be tested for lipid phosphatase activity, as an alternative to overexpression and purification from insect cells or yeast. To address this question, myotubularin and PTEN were transiently over-

FIG. 3. Dephosphorylation of fluorescent phosphoinositide substrates by MTM1 and PTEN immunoprecipitates. FLAG epitopetagged MTM1 and PTEN wild-type and Cys to Ser mutant proteins were transiently overexpressed in Cos 1 cells and purified by immunoprecipitation with anti-FLAG affinity resin. (A) Wild-type (right) and C375S (left) myotubularin immunoprecipitates were incubated with 1.5 ␮g di-C 6-NBD6-phosphatidylinositol 3-phosphate for 30 min at 30°C, processed as described under Materials and Methods, and analyzed by TLC. The origin, solvent front, and migration positions of authentic di-C 6-NBD6-phosphoinositide standards are shown (at right). The di-C 6-NBD6-phosphatidylinositol reaction product generated by wild-type MTM1 is indicated by a (white arrow). (B) Wild-type (right) and C124S (left) PTEN immunoprecipitates were incubated with 1.5 ␮g di-C 16-NBD6-phosphatidylinositol 3,4,5trisphosphate for 30 min at 37°C, processed as described under Materials and Methods, and analyzed by TLC. The origin, solvent front, and migration positions of authentic di-C 6-NBD6-phosphoinositide standards are shown (at right). The di-C 16-NBD6-phosphatidylinositol 4,5-bisphosphate reaction product generated by wild-type PTEN is indicated by a (white arrow).

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shown). Collectively, these results demonstrate that this methodology is useful for testing the lipid phosphatase activity of proteins expressed exogenously and immunoprecipitated from mammalian cells. In summary, we have provided evidence that fluorescent phosphoinositides can be effectively utilized to examine the activity and specificity of inositol lipid phosphatases. The procedure outlined here is simple, rapid, and can be used with recombinant proteins expressed in bacteria or immunoprecipitated from mammalian cells. Because quantitation of phosphatase activity by this method is only approximate, it is most useful for the determination of enzyme substrate specificity. As such, it effectively complements the malachite green-based assay procedure reported previously, which is most useful for the accurate quantitation of lipid phosphatase activity where the substrate specificity is known (1). Acknowledgments. This work was supported by a grant from the National Institutes of Health and the Walther Cancer Institute. G.T. is supported by a postdoctoral fellowship from the Endocrinology and

Metabolism Training Grant, Michigan Diabetes Research Training Center.

REFERENCES 1. Maehama, T., Taylor, G. S., Slama, J. T., and Dixon, J. E. (2000) A sensitive assay for phosphoinositide phosphatases. Anal. Biochem. 279, 248 –250. 2. Maehama, T., and Dixon, J. E. (1998) The tumor suppressor, PTEN/MMAC1, dephosphorylates the lipid second messenger, phosphatidylinositol 3,4,5-trisphosphate. J. Biol. Chem. 273, 13375–13378. 3. Taylor, G. S., Maehama, T., and Dixon, J. E. (2000) Myotubularin, a protein tyrosine phosphatase mutated in myotubular myopathy, dephosphorylates the lipid second messenger, phosphatidylinositol 3-phosphate. Proc. Natl. Acad. Sci. USA 97, 8910 – 8915. 4. Okada, T., Hazeki, O., Ul, M., and Katada, T. (1996) Synergistic activation of PtdIns 3-kinase by tyrosine-phosphorylated peptide and ␤␥-subunits of GTP-binding proteins. Biochem. J. 317, 475– 480. 5. Blondeau, F., Laporte, J., Bodin, S., Superti-Furga, G., Payrastre, B., and Mandel, J. (2000) Myotubularin, a phosphatase deficient in myotubular myopathy, acts on phosphatidylinositol 3-kinase and phosphatidylinositol 3-phosphate pathway. Hum. Mol. Genet. 9, 2223–2229.