Supplemental Information s1

SUPPLEMENTAL INFORMATION

MATERIALS AND METHODS

Materials. Bodipy TMR-PtdIns(4,5)P2-C16, Glo-( Bodipy-TMR-PtdIns(4,5)P2-C16),

Bodipy TMR-HyPl(4,5)P2, and Bodipy TMR-HyPl(3,4)P2 were purchased from Echelon

Bioscience (Salt Lake City, UT). The first fluorescent PIP2 species has the label on a chain and regular ester linkage of the two chains to the glycerol backbone (no longer commercially available); the second also has the label on a chain, but an amide linkage of the chain to the backbone (Echelon catalog number C-45M16a); the third has two C16 chains plus the label (Echelon catalog number H-45TM). Gambhir et al., (2004) shows the structure of the first species; structures for the remaining species can be viewed on the

Echelon Web site (www.echelon-inc.com). We obtained identical (within experimental error) results with all three species in FCS measurements and pooled the results. We stress that all fluorescent PIP2 molecules had long chains and partitioned strongly into bilayers. 1,2-Dioleoyl-sn-Glycero-3-Phosphoethanolamine-N-(Lissamine Rhodamine B

Sulfonyl), 1-Arachidoyl-2-Hydroxy-sn-Glycero-3-Phosphocholine (20- carbon chain lyso-PC), and 1-Oleoyl-2-[12-[[6-[(7-nitrobenz-2-oxa-1,3-diazol-4- yl)amino]hexanoyl]amino]-dodecanoyl]-sn-Glycero-3-Phosphoinositol-4,5-trisphosphate

(NBD-PIP2) were purchased from Avanti Polar Lipids (Alabaster, AL). 1,1’- dioctadecyl-3,3,3’,3’-tetramethylindodicarbocyanine, as a 4-chlorobenzenesulfonate salt

(DiD; also known as DiI-C18) was from Molecular Probes (Eugene, OR). Rhodamine B, used to calibrate the width of the confocal beam, was from Sigma (St. Louis, MO). We

1 used both commercially available needles from Eppendorf and needles we pulled ourselves for the microinjection experiments. Thin-wall single-barrel standard borosilicate glass tubes (with filament), outer diameter 1.0 mm and inner diameter 0.75 mm, were from World Precision Instruments (Sarasota, FL). Needles were pulled on a

Flaming Brown micropipette puller, model P.80/PC, Sutter Instrument Co (Novato, CA).

Because the tip diameter of these needles was slightly larger, we had fewer problems with micelle aggregation clogging the tips. We used 35 mm glass bottom dishes from

MatTek Corporation (Ashland, MA) to hold the cells during microinjection and FCS measurements.

Cell culture. Except where noted, all cells were maintained at 37°C in a 5% CO2 environment. NIH 3T3, HeLa, and HEK293 cells were grown in Dulbecco’s modified

Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 50 units/ml of penicillin, and 50 g/ml streptomycin sulfate. COS1 cells were grown in DMEM supplemented with 8% FBS, 50 units/ml of penicillin, and 50 g/ml streptomycin sulfate.

REF52 cells were grown in DMEM medium supplemented with 10% FBS, 50 units/ml of penicillin, and 50 g/ml streptomycin sulfate; the 37°C environment contained 6.3%

CO2. We typically starved the cells in serum-free medium for 1 – 2 hours prior to an experiment. Where specified, cells were starved in serum-free medium for periods ranging from 4 hours to overnight.

Sample preparation. We injected three different forms of Bodipy TMR-PIP2 micelles.

2 1) Bodipy TMR-PIP2 micelles were formed by dissolving the fluorescent PIP2 in 1 mM

EDTA (pH 7). Centrifugation of the sample before filling the needle helped to remove large aggregates. The main difficulty was aggregation of micelles in the needle tip, presumably because of exposure to calcium ions (~1 mM) in the bathing solution (Hirai et al., 1996; Flanagan et al., 1997). Standard methods for cleaning the needles (applying high pressure to blow the solution from a needle placed far from the cells; moving the needle rapidly in and out of the medium) often were ineffective, necessitating frequent needle replacement. The needle clogging problems caused many experimental failures and we investigated other ways of delivering PIP2.

2) Unlabeled PIP2 / Bodipy TMR-PIP2 micelles (4:1 ratio). Because PIP2 micelles are somewhat larger than Bodipy TMR-PIP2 micelles (FCS measurements, not shown), we hoped they would be more stable. We again encountered significant problems with aggregation of micelles inside the needle and on the needle tip. We tried to inject cells in a bathing medium with reduced calcium (100 μM), but cell survival was very poor under these conditions. This led us to investigate forming micelles of fluorescent PIP2 with lyso-PC, which aggregate less markedly when exposed to ~ 1 mM Ca2+ in the bathing solution.

3) Arachidoyl-lysoPC / Bodipy TMR-PIP2 micelles were formed from a mixture of 82% lysoPC and 18% Bodipy TMR-PIP2. We added a 1 mM EDTA (pH = 7.0) solution to dried lipids to form a micelle suspension, mixed the suspension, warmed it to 37ºC, diluted it to a total lipid concentration of 500 M with 1 mM EDTA, briefly (~5 s)

3 sonicated it in a bath sonicator, then added water to produce the desired concentration for microinjection (~ 125 – 250 M for all types of micelles injected). The solution was warmed to 37ºC before filling the needles. The main difficulty, again, was aggregation.

We used standard methods to clean the needle when the micelles aggregated inside the needle and on the needle tip: the microinjector’s “CLEAN” function (blowing needle with high pressure into the medium far away from cells), its “HOME” function (moving needle rapidly out and back in to the solution), mock injections into the medium in an area far away from cells, varying injection pressure, and changing needles. Injection of these micelles caused one or a few large blebs to form in some cells. We do not understand the phenomenon, but note it occurred most often when the microinjection site was close to the cell nucleus. We took advantage of bleb formation by measuring PIP2 diffusion in these regions, which presumably lack many cytoskeletal proteins, and comparing these values of D with those obtained on native unperturbed Rat1 plasma membranes.

We injected rhodamine-PE either in ethanol solution or in the form of arachidoyl-lysoPC micelles (30% rhodamine-PE, 70% lysoPC) prepared as described above. Cell survival was poor following injections of the ethanol solution because removal of the needle often damaged the plasma membrane. We measured similar values of D with both rhodamine-

PE solutions, however, and pooled the data.

Microinjection. Cells were grown on glass bottom dishes for two days to achieve 60 –

80% confluence at the time of microinjections. We used an InjectMan NI2 with

4 FemtoJet pump from Eppendorf to microinject the solutions into cytoplasm. We typically set the injection pressure Pi = 17 – 25 hPa, and kept the compensation pressure

Pc = 0 hPa to avoid leakage of the sample from the needle. Injection time usually was set at t = 0.5 s, but ranged from t = 0.2 – 0.6 s. Typically, we injected about 10-25 cells within a 10 – 20 minute period. We examined the microinjected cells under the phase microscope (Axiovert 200M from Zeiss with 40 × phase 2 objective) to select viable cells that had an appropriate level of fluorescence in the plasma membrane for FCS measurements. (We usually observed between 1 and 100 fluorescent PIP2 in the confocal volume, and the relative fractions of fluorescent PIP2 in the plasma membrane and the cytoplasm shown in Figure 1C was typical of the hundreds of cells we measured. When blebs formed, we observed a higher ratio of membrane:cytoplasmic PIP2.) We then changed the medium to phenol-free Leibovitz’s-15 (L-15), transferred the cells to the

FCS microscope and monitored fluorescence by scanning through individual cells (Figure

1C). The FCS measurements of diffusion of fluorescent PIP2 in the plasma membrane typically occurred 20 – 30 min after microinjections. We tested the effect of time between microinjection and FCS measurements by injecting cells on the FCS microscope and performing measurements ≤2 minutes after delivery of fluorescent PIP2; we also measured PIP2 diffusion in cells incubated at 37 ºC for ~45 min in full growth medium.

We measured the D of fluorescent PIP2 on the outer leaflet of the plasma membrane by adding a solution of micelles (all 3 types of PIP2 micelles) to the cells, incubating the dish at room temperature for ~10 min, and rinsing the cells with phenol-free medium three times before imaging the cells on the FCS microscope. In experiments using DiD, adding

5 the lipid to the bathing solution induced endocytosis in some cells. We did not collect data from the cells that either had fluorescent lipids in internal organelles or had fluorescent (presumably endocytotic) vesicles in the cytoplasm.

The measurements on GUVs shown in Figure 3 used GUVS formed from 9:1

POPC/POPS using the gentle hydration method described in detail elsewhere

(Golebiewska et al., 2006). The bathing solution contained 100 mM KCl, 10 mM

MOPS, pH 7.0. We then exposed the GUVS to lysoPC/Bodipy TMR-PIP2 (5:1) micelles in the bathing solution. Previous measurements showed the D of Bodipy TMR-PIP2 in

GUVS was the same when PIP2 was either transferred from exposure to micelles or included in the lipid mixture used to form the vesicles.

FCS measurements. Confocal imaging and FCS measurements were performed on a

Zeiss LSM 510 Meta/Confocor 2 apparatus (Jena, Germany) using standard configurations. We selected minimal laser powers to avoid photobleaching of the fluorescent probes. We used a 40 × NA 1.2 C-Apochromat water immersion objective and adjusted pinholes at least daily. We excited Bodipy TMR and rhodamine with the

543 nm HeNe laser and collected emission spectra through a 560 LP filter.

We calibrated the detection volume by measuring the diffusion of rhodamine (RhB, D =

3 × 10-6 cm2/s) in water (Thompson, 1991; Hess and Webb, 2002). The 1/e2 radius of the detection volume for the 543 nm line was r = 0.19 ± 0.01 μm. We monitored the count rate during data acquisition and rejected measurements that were visibly higher or lower than normal to avoid artifacts due to bleaching and/or cell movement. We used Sigma

6 Plot and a least squares algorithm to fit the autocorrelation curves to the model equation for free Brownian diffusion in two dimensions commonly used in FCS (Schwille et al.,

1999):

1 1 G( )    N 1  S1  d

where τd is the average residence time and N is the average number of fluorescent PIP2 in the measurement volume. We analyzed cells that had N = 1 – 100, which corresponds to fluorescent PIP2 comprising 0.001% – 0.1% of lipids in the plasma membrane (the

5 confocal volume contains ~ 10 lipids). Thus, the level of fluorescent PIP2 is negligible compared to the endogenous level of unlabeled PIP2.

We calculated the diffusion coefficient, D, from the Einstein relation:

r 2 D  S2 4 d

If the measured volume contains different populations of molecules with the same fluorescence quantum yields (e.g. free fluorescent probe diffusing in the same volume as fluorescent PIP2) the equation becomes:

1 Y G( )   i N   i 1  S3  d ,i

7 where N is number of molecules, Yi is a fraction of molecules diffusing with diffusion

2 coefficient Di producing residence times τd,i = r /4Di. For the rhodamine PE measurements we observed ~20% free probe or autofluorescence, which diffused more rapidly than the membrane-bound rhodamine PE. With Bodipy TMR-PIP2, we observed only a negligible (typically <10%) fraction of rapidly diffusing fluorescence. All measurements were performed at room temperature, 25 ± 1ºC, monitored throughout the experiment using a thermocouple.

In some cells (< 5% for the Bodipy TMR-PIP2 micelles; never for the Lyso PC –Bodipy

TMR-PIP2 micelles) a significant fraction of the PIP2 in the confocal volume appeared to be bound irreversibly. The fraction is possibly due to: micelles that adsorbed to the cytoskeleton, but did not incorporate PIP2 into the phospholipid leaflet; PIP2 inclosed in

“fences” that form compartments smaller than the confocal area; PIP2 adsorbed irreversibly to membrane proteins or that dissociates much more slowly than the time for diffusion out of the confocal area (~10 ms). For measurements on these cells, we omitted the first 10 s of fluctuation recording time from the record: the irreversibly bound Bodipy

TMR-PIP2 was bleached during this time, and diffusion of the fluorescent PIP2 in the plasma membrane could be recorded from analysis of subsequent 10 second records of fluorescence fluctuation.

Statistical Analysis. We used software provided by Zeiss and SigmaPlot (SPSS, INC.,

Chicago, IL) for curve fitting and SigmaStat (SPSS, INC., Chicago, IL) for statistical

8 analysis. We compared the values of diffusion coefficient of fluorescent lipids using

Kruskal-Wallis One Way Analysis of Variance on Ranks, Dunn’s Method, One Way

Analysis of Variance, Tukey’s Method, and paired t-test (Zar, 1998). We concluded the values are significantly different when P < 0.05.

THEORETICAL ANALYSIS

Analysis of PIP2 diffusion and simultaneous rapidly reversible binding. Our analysis follows closely the simple derivation in Crank’s “The Mathematics of

Diffusion”, Chapter 14 (Crank, 1975). Reports by Elson (Elson and Reidler, 1979;

Icenogle and Elson, 1983), Koppel, (1981), and Jacobson et al., (1984) may be consulted for a more extensive discussion and analysis of this simultaneous binding/diffusion problem. If the diffusion of PIP2 in the two dimensional plasma membrane located in the x-y plane is accompanied by reversible adsorption, the diffusion equation (Fick’s second law) becomes

C   2C  2C  S  D     free  2 2  S4 t  x y  t

where C is concentration of PIP2 free to diffuse, S is concentration of sequestered PIP2

(PIP2 reversibly bound or immobilized), and Dfree is the diffusion coefficient of the free

PIP2. In the simplest case, S is linearly proportional to C ( S  RC ) as predicted by

Henry’s law, to which all adsorption isotherms reduce at low concentration.

Combination of these two equations produces a diffusion equation with an apparent

9 diffusion constant, D = Dfree/(R + 1). In other words, reversible association scales the diffusion coefficient by the factor 1/(R+1):

C D   2C  2C   free     2 2  S5 t R  1  x y 

If T = S + C, where T = total concentration of PIP2, then C = T/(R +1). Thus the concentration of free PIP2 is related to the total concentration by the same scale factor of

1/(R + 1).

This derivation assumes sequestration occurs rapidly with respect to diffusion. The available experimental evidence suggests that the electrostatic association and dissociation of PIP2 from an unstructured cluster of basic residues does indeed occur rapidly compared to the time for a FCS diffusion measurement of PIP2 in a plasma membrane (about 10 ms, Figure 2). Specifically, the experimentally or theoretically determined electrostatic energy of interaction of a single PIP2 with a basic cluster on a peptide corresponding to the MARCKS effector domain (or Lys13 or Lys7) is about 3 kcal/mol (Wang et al., 2002; Wang et al., 2004); the equilibrium association constant is

K ~ 102 M-1. The forward rate constant, estimated from stop flow measurements of the binding of basic peptides to membranes containing PIP2, is about diffusion limited or kon

6 -1 -1 4 -1 ~ 10 M s . Thus, the dissociation rate constant is kd ~ 10 s , or the lifetime of a PIP2 in the electrostatic well adjacent to a basic cluster is of order ~ 0.1 ms, significantly less than the diffusion time of 10 ms. In other words, our diffusion measurements suggest an individual PIP2 diffusing out of the confocal volume in a typical time of 10 ms, spends

10 2/3 of its time (7 ms) in electrostatic wells, and 1/3 of its time (3 ms) in free diffusion.

The average time spent in a single electrostatic well is only 0.1 ms. The PIP2 presumably encounters many basic clusters during its random diffusional walk across the confocal area (r ~200 nm). This area of the inner leaflet contains ~ 105 phospholipids, ~ 3 × 103

2 3 PIP2 and ~ 10 - 10 MARCKS proteins, so our assumption a PIP2 encounters many clusters of basic residues while diffusing in this area is reasonable.

RESULTS

Exposing Large Unilamellar Vesicles (LUVs) to NBD-PIP2 micelles or lysoPC-NBD-

PIP2 mixed micelles transfers PIP2 only to the outer leaflet. We exposed 100 nm diameter extruded PC LUVs to NBD-PIP2 micelles or lysoPC-NBD-PIP2 mixed micelles and monitored the fluorescence signal. Addition of LUVs to NBD-PIP2 micelles produces a 5-fold increase in fluorescence due to the release of self-quenching. Adding the membrane-impermeable quencher sodium dithionite reduced fluorescence to 1% of the initial value. When we formed the PC/PIP2 vesicles from a mixture of both lipids, i.e.

NBD-PIP2 was equally distributed on both leaflets, adding sodium dithionite reduced the fluorescent signal only ~50%. These control experiments support our hypothesis that the micelles deliver PIP2 to only the exposed leaflet of a LUV, and thus our assumption injecting PIP2 micelles into cells delivers the fluorescent lipid only to the inner leaflet of the plasma membrane. Transfer of lipids into membranes is discussed in detail elsewhere

(Pagano et al., 1981; Elvington and Nichols, 2007).

11 Why do the PIP2 micelles deliver PIP2 to the plasma membrane rather than internal membranes? Figure 1C suggests that a major fraction of the PIP2 micelles injected into a cell transfer PIP2 into the plasma membrane (PM) rather than internal membranes. We wondered why this happened. Internal membranes have a larger surface area and a less negative surface potential, and one might expect the highly negatively charged micelles would deliver the PIP2 more effectively to the internal membranes. We tested this expectation with model phospholipid vesicles: we mixed sucrose-loaded 3:1 PC/PS

LUVs with an equal number of PC LUVs and added PIP2 micelles, then centrifuged the solution to separate the vesicles. When both types of LUVs were present, >90% of the fluorescence remained in the supernatant, i.e. the micelles transferred PIP2 mainly into the electrically neutral PC vesicles, as expected. If the PC vesicles were omitted from the mixture, > 90% of the fluorescence was in the pellet of sucrose-loaded PC/PS vesicles; i.e. the micelles transferred PIP2 into the PC/PS vesicles.

We considered one possible explanation for the preferential incorporation of PIP2 into the plasma membrane in our experiments: this membrane contains a relatively high concentration of proteins with membrane-bound basic clusters (e.g. MARCKS). The injected PIP2 micelles diffusing randomly through the cell could adsorb to these basic clusters and thus deliver the PIP2 monomers preferentially to the plasma membrane. We tested this possibility by adding a basic peptide, MARCKS(151-175), to the mixture of

PC and sucrose-loaded PC/PS vesicles (10 μM accessible lipid in the form of 3:1 PC/PS vesicles, 10 μM accessible lipid in the form of PC vesicles, 200 nM MARCKS(151-175) in 100 mM KCl, 10 mM MOPS, pH 7). MARCKS(151-175) binds preferentially to the

12 PC/PS vesicles (Arbuzova et al., 2000; Rusu et al., 2004). Adding PIP2 micelles (or lyso-

PC/PIP2 micelles) and centrifuging the mixtures showed that PC/PS vesicles with bound

MARCKS peptide incorporate a significant faction of PIP2 from the micelles; ~ 50% of the fluorescence was associated with the pellet fraction containing these vesicles. The result is consistent with the postulate that the PIP2 micelles deliver PIP2 preferentially to the plasma membrane because of membrane-bound clusters of basic residues, but does not rule out other interpretations.

Exposure of cells to amphipathic weak bases, Ca2+ ionophore or carbachol increases the D of PIP2. If PIP2 is bound to clusters of basic residues, it should be released when amphipathic weak bases bind to the inner leaflet and reverse its net negative charge, which will produce desorption of the basic clusters. For example, 2 M sphingosine reverses the charge on a PC/PS vesicle, and causes basic peptides to desorb from the phospholipid vesicles (McLaughlin et al., 2005); it might also release clusters of basic residues from the inner leaflet of a plasma membrane, and thus release sequestered PIP2.

We tested this by measuring the D of Bodipy TMR PIP2 before and after adding 2 M sphingosine to Cos1 cells: we observed a small but statistically significant increase, from

0.9 ± 0.1 m2/s to 1.2 ± 0.3 m2/s (± SD, n = 9). The amphipathic weak base trifluoperazine (TFP) also reduces the net negative charge on a PC/PS vesicle and causes desoprtion of basic peptides (Sengupta et al., 2007). Adding 25 M TFP to Rat1 cells also produced a small, statistically significant increase in D, from 1 ± 0.2 m2/s to 1.3 ±

0.2 m2/s (± SD, n = 17). We observed similar effects after adding the G protein-coupled

13 receptor (muscarinic and nicotinic) agonist carbachol and the calcium ionophore A23187, but these experiments should be regarded as preliminary and are difficult to interpret for several reasons. The biologically important question, in our opinion, is whether local increases in Ca2+, and thus calcium/Calmodulin (Ca/CaM), produce local increases in free

2+ PIP2. There is evidence that localized increases in Ca should be more effective than global increases in Ca2+ (in the entire cytoplasm) in increasing Ca/CaM, and perhaps

2+ consequently free PIP2. Specifically, when the global concentration of Ca increases, the free concentration of Ca/CaM declines ~100-fold rapidly (few seconds) (Black et al.,

2004); this is because many proteins in the cell bind Ca/CaM with high affinity.

Detecting and measuring local changes in the free concentration of PIP2 in the membrane in response to local increases in Ca2+ will, of course, be more difficult, but such experiments could provide important information and remain a challenge for the future.

Measurements of the partitioning of GFP-PH domains between the plasma membrane and the cytoplasm are consistent with our conclusion that 2/3 of the PIP2 is bound reversibly. Consider a hypothetical spherical cell of diameter 10 μm and assume its plasma membrane has the same mole fraction of PIP2 as a human erythrocyte, i.e. PIP2 comprises ~3% of phospholipids on the inner leaflet (Ferrell and Huestis, 1984;

Christensen, 1986; Hagelberg and Allan, 1990). If the membrane has 1 phospholipid/100

2 2 Å , there are ~30,000 PIP2 per μm . (The data for PIP2 in cells other than erythrocytes are less reliable, as discussed by Hilgemann, (2007); his more detailed estimates suggest that for mammalian cell lines (e.g. HEK293, CHO) and cardiac tissue the PIP2 density in the surface membrane is “minimally 20,000 and possibly as high as 60,000 per μm2”.) The

14 equivalent total concentration in our hypothetical cell is T = 30 μM PIP2 (total PIP2 concentration assuming it is dissolved uniformly in the interior of the cell), and our diffusion measurements suggest the sequestered concentration of PIP2 is S = 20 μM, and its free concentration is C = 10 μM.

Our conclusion that C = 10 μM is reasonable in the context of experiments examining the partitioning of the GFP constructs of the PLC-δ1 PH domain (Kd = 2 μM for the 1:1 complex with PIP2 in model membranes) and pleckstrin (Kd = 30 μM) between cytoplasm and the plasma membrane in a typical cell. Extensive measurements on many cell types show most of the GFP-PLC-δ1 PH domain is bound to the plasma membrane (Stauffer et al., 1998; Varnai and Balla, 1998; Kwik et al., 2003; Varnai and Balla, 2006), which suggests the free concentration of PIP2 is > 2 μM. Most of the pleckstrin PH domain, on the other hand, is free in the cytoplasm (Lemmon and Ferguson, 2000), which suggests the free concentration of PIP2 is < 30 μM. These estimates of the free PIP2 level with PH domains are not without complications (Varnai and Balla, 2006), but the simplest interpretation is that 2 μM < C < 30 μM, and this is consistent with our conclusion that about 10 μM PIP2 is free in the plasma membrane and capable of binding these PH domains.

Non-uniform PIP2 distribution in cells. In certain cells proteins with clusters of basic residues are concentrated in specific regions. For example, MARCKS is present at high concentrations in ruffles (Myat et al., 1997; Myat et al., 1998) and nascent phagosomes

(Allen and Aderem, 1995); GAP43 in neural growth cones (Skene, 1990); and syntaxin in

15 regions of cells where exocytosis occurs (Sieber et al., 2007). Peptides corresponding to the basic clusters on MARCKS (Gambhir et al., 2004), GAP43 and the juxtamembrane region of syntaxin (FRET data, not shown, similar to data in Gambhir et al., (2004)) laterally sequester PIP2. This suggests there should be higher mole fractions of PIP2 in ruffles (Honda et al., 1999), nascent phagosomes (Botelho et al., 2000), the growth cone of neurons (Laux et al., 2000) and regions of cells where exocytosis occurs (Milosevic et al., 2005; Aoyagi et al., 2005). For example, recent measurements suggest cells have regions containing 75 syntaxin/400 nm2 (see Figure 4C of Sieber et al., (2007)). The basic juxtamembrane region of syntaxin is localized to the membrane (Kim et al., 2002) and peptides corresponding to this region (KKAVKYQSKARRKK, net charge +8) sequester ~2 PIP2. This would mean such a 20 nm square region would contain 150 PIP2, comprising 30% of the membrane phospholipids.

The high concentration of basic clusters in specific regions of the plasma membrane should not only concentrate PIP2, but also reduce the local diffusion coefficient of PIP2.

This will allow the PIP-kinases that are concentrated with MARCKS in ruffles and nascent phagosomes (Doughman et al., 2003), and with syntaxin in regions undergoing exocytosis in PC12 cells (Aoyagi et al., 2005) to function more effectively to increase the local free concentration of PIP2. Specifically, if the diffusion coefficient were not significantly reduced, PIP2 would rapidly diffuse away from regions of local synthesis.

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