A High Avidity Biosensor Reveals PI(3,4)P2 is Predominantly a Class I PI3K Signaling Product

Class I PI 3-kinase (PI3K) signaling is central to animal growth and metabolism, and disruption of this pathway occurs frequently in cancer and diabetes. However, the specific spatial/temporal dynamics and signaling roles of its minor lipid messenger, phosphatidylinositol (3,4)-bisphosphate [PI(3,4)P2], are not well understood. This owes principally to a lack of tools to study this scarce lipid. Here, we developed a high sensitivity genetically encoded biosensor for PI(3,4)P2, demonstrating high selectivity and specificity of the sensor for the lipid. We show that despite clear evidence for class II PI3K in PI(3,4)P2-driven function, the overwhelming majority of the lipid accumulates through degradation of class I PI3K-produced PIP3. However, we show that PI(3,4)P2 is also subject to hydrolysis by the tumor suppressor lipid phosphatase PTEN. Collectively, our results show that PI(3,4)P2 is potentially an important driver of class I PI3K-driven signaling, and provides powerful new tools to begin to resolve the biological functions of this lipid downstream of class I and II PI3K.


Introduction
Class I phosphoinositide 3-OH kinase (PI3K) signaling is central to the control of growth and metabolism in animals (Vanhaesebroeck et al., 2012). Overactivation of this pathway is the most common event in cancer (Fruman et al., 2017), yet given its major role in insulin signaling, inhibition of the pathway triggers insulin resistance and type 2 diabetes (Hopkins et al., 2018). Therefore, the ability to selectively manipulate PI3K signaling could have tremendous therapeutic benefit. Efforts to accomplish this goal are a major focus of the biomedical enterprise (Fruman et al., 2017) At the molecular level, PI3K signaling involves the generation of the plasma membrane (PM) second messenger lipids phosphatidylinositol 3,4,5-trisphosphate (PIP3) and phosphatidylinositol 3,4-bisphosphate [PI(3,4)P2] that activate downstream effector proteins like the serine/threonine kinase Akt. PIP3 is the major lipid produced, and most functions of the pathway are attributable to it (Vanhaesebroeck et al., 2012). PI(3,4)P2 has instead been viewed as either a degradation product (Ishihara et al., 1999), or as an alternative activator of the pathway (Ebner et al., 2017). However, selective functions for PI(3,4)P2 have recently been described that are independent of PIP3 (Li and Marshall, 2015). These include the formation of lamellipodia and invadopodia (Krause et al., 2004;Bae et al., 2010;Oikawa et al., 2008;Sharma et al., 2013), along with clathrin-mediated and clathrin-independent endocytosis (Posor et al., 2013;Boucrot et al., 2015) In each case, these functions could conceivably be driven by, or occur independently of, class I PI3K signaling.
Resolving how the spatial/temporal dynamics of PI(3,4)P2 signaling couples to different biological functions requires approaches to identify the lipid in intact, living cells. Isolated lipid binding domains fused to fluorescent reporters often make highly selective genetically encoded biosensors for this purpose . The pleckstrin homology (PH) domain on the Cterminus of TAPP1 (TAndem Ph-domain containing Protein 1) exhibits specific binding to PI(3,4)P2 in the test tube (Thomas et al., 2001;Dowler et al., 2000). As a result, several studies have employed fluorescent protein conjugates of this domain to track PI(3,4)P2 signaling, though the domain fails to detect resting levels or the limited accumulation of the lipid in response to stimuli such as insulin-like growth factor (Kimber et al., 2002;Marshall et al., 2002;Oikawa et al., 2008;Posor et al., 2013).
Herein, we developed a higher-avidity tandem trimer of PH-TAPP1. We show PI(3,4)P2 generation is sufficient to recruit the probe, which is exquisitely selective for the lipid over other phosphoinositides. We then demonstrate that the class I PI3K pathway, acting via PIP3 synthesis, dominates PI(3,4)P2 accumulation in cells. Our data also support the recently proposed direct degradation of both PI(3,4)P2 and PIP3 by the lipid phosphatase and tumor suppressor PTEN (Malek et al., 2017). Collectively, our data show that the class I PI3K pathway is the most potent driver of PI(3,4)P2-dependent signaling.
Previous studies using the isolated TAPP1 C-terminal PH domain (cPH) as a lipid biosensor found no detectable translocation in response to stimuli that induce modest PI(3,4)P2 accumulation (Kimber et al., 2002). To improve avidity, tandem dimers of TAPP1 fragments containing the cPH domain have been employed (Oikawa et al., 2008;Posor et al., 2013;He et al., 2017). However, these constructs are based on a fragment including the entire C-terminus (Furutani et al., 2006), which carries binding sites for other proteins in its tail, and have been shown to induce dominant negative effects (Kimber et al., 2002;Hogan et al., 2004;Thalappilly et al., 2008;Li and Marshall, 2015). Therefore, we built tandem dimers and trimers of the isolated cPH domain (residues 169-329 in human TAPP1) as previously defined (Marshall et al., 2002), and included a nuclear export sequence (figure 1A).
Expression of EGFP-tagged cPH monomers, dimers or trimers in COS-7 cells maintained in 10% serum exhibit a predominantly cytosolic localization of the probe when viewed by confocal microscopy ( figure 1B). However, comparison with another cytosolic protein (infra-red fluorescent protein, iRFP) revealed enrichment of cPHx2 and cPHx3 at the cell periphery; normalization of the two signals followed by subtracting the iRFP signal from that of EGFP (see Materials and Methods) showed striking cPHx2/3 peripheral localization (figure 1B). This signal was abolished by a PM-targeted INPP4B phosphatase that specifically degrades PI(3,4)P2 (Gewinner et al., 2009), though not by a catalytically inactive mutant (figure 1B). Quantification of the difference between EGFP and iRFP signals (figure 1C with Kruskal-Wallis statistic 152.5; p < 10 -4 ) yielded significantly increasing signal from cPHx1 to x3; INPP4B-CAAX significantly reduced cPHx3 signal (p < 10 -4 in each case, Dunn's multiple comparison test), whereas the C842A inactive mutant did not (p > 0.99). Therefore, cPHx2 and cPHx3 could detect PI(3,4)P2 at the plasma membrane of living cells in the presence of serum.
A potential caveat to using tandem arrays of lipid binding domains is that the resulting probe may cluster lipids and exhibit aberrant localization or mobility in the membrane. To determine whether this was the case, we performed single molecule imaging of cPHx1, x2 and x3 mobility in the plasma membrane by tagging with photoactivatable mCherry (PAmCherry). As shown in figure 1D, full activation of the probe with violet light led to uniform labelling of the plasma membrane in total internal reflection fluorescence microscopy (TIRFM), whereas low activation intensities allowed us to image single molecule trajectories (Manley et al., 2008). Analysis of the mean square displacement of these trajectories over time revealed free Brownian motion of all three probes ( figure 1E). cPHx1, x2 and x3 diffused with an apparent diffusion coefficient of ~0.3 µm 2 /s (figure 1F) and did not substantially differ (F = 0.71, p = 0.50, one-way ANOVA), and all exhibited a relative deviation ratio close to 1 (figure 1G), meaning there was no substantial difference in actual displacement relative to that predicted by free Brownian diffusion (Fujiwara et al., 2016). Therefore, the tandem array of domains did not slow the probe's mobility in the membrane.
The lifetime distribution of the single molecules on the membrane followed a single exponential decay with a characteristic "off rate" constant (koff) that increased linearly with laser power due to increased photobleaching ( figure 1H). Extrapolation of this off rate to the intercept at zero illumination power allowed us to estimate the true off rate of the probes. cPHx1 has a lifetime on the membrane of approximately 90 ms (off rate = 11 s -1 ), whereas cPHx2 and cPHx3 both showed lifetimes of approximately 140 ms. This change in dissociation rate of the tandem arrays was comparatively modest, and certainly not multiplicative as would be expected from multi-ligand binding. This implies that, for the majority of probe molecules at steady state, the PH domains are not all occupied by lipid, most likely because of the low abundance of PI(3,4)P2.
The enhanced PM binding observed in figure 1C likely stems from transient dual or triple occupancy of the PH domains slowing the overall off-rate of the population (and perhaps enhancing the on rate for cPHx3 vs cPHx2).
A potential limitation of biosensors is that when bound to lipid, they will protect the lipid from consumption by enzymes and occlude interaction with endogenous effector proteins, both of which may be needed for local enrichment of the lipid. Therefore, free diffusion of the probe:lipid complex can disrupt local enrichment rather than reporting on it. From the relationship r = (2D/koff) 0.5 (Teruel and Meyer, 2000), the distance, r, that cPHx3 typically diffuses while attached to lipid can be estimated as ~290 nm, not much larger than the diffraction limit. This implies there will be a limited propensity of the probes' free diffusion to "smear out" local enrichment of PI(3,4)P2 molecules that are detectable with diffraction-limited optical imaging.
Given the much higher avidity of cPHx3 and x2 for the PM, we wanted to rule out a weak interaction with other lipids that might synergize to influence PM targeting by these probes. To this end, we used chemically induced dimerization again, this time using the INPP5E phosphatase domain (figure 1L), since this domain is active against PI(4,5)P2 and PIP3, generating PI4P and PI(3,4)P2, respectively (Asano et al., 1999;Kong et al., 2000). INPP5E recruitment did not displace PI4P-binding P4Mx1 or any of the cPH probes from the PM, though it did displace probes that bound to PIP3 or PI(4,5)P2 (figure 1M). Therefore, these two inositol lipids do not influence the PM-association of cPH probes.
Together, these results allowed us to conclude that (i) the cPHx3 probe derived from TAPP1 exhibits significantly greater localization than previously used single and tandem dimer versions, (ii) the tandem trimer configuration does not disrupt free diffusion in the plane of the membrane nor sequester PI(3,4)P2 to an extent that prevents INPP4B access and (iii) exhibits localization that depends solely on the phosphoinositide PI(3,4)P2. These are crucial criteria in the definition of a reliable genetically encoded lipid biosensor. However, there is another critical criterion that must be met, which we tested next.

PI(3,4)P2 is sufficient for cPHx3 cellular localization
We have argued that a crucial but often ignored criterion for a high-fidelity lipid biosensor is a sole requirement of the lipid to drive biosensor localization . A convenient way to test this criterion is to induce ectopic synthesis of the lipid elsewhere in the cell and test whether this is sufficient to recruit the biosensor (Hammond et al., 2014). To this end, we turned to the Legionella effector protein LepB, the N-terminus of which possesses PI3P 4-OH kinase activity, thus generating PI(3,4)P2 via a non-canonical pathway not utilized by mammalian cells (Dong et al., 2016). We found that expression of LepB led to substantial PI3P depletion from cells, and consequent swelling of the endosomal compartment that depends on this lipid for function (B.D.G and G.R.V.H., unpublished observations). Therefore, to induce acute LepB activity, we turned to an optogenetic approach that utilizes genetic code expansion to incorporate an unnatural, caged amino acid into the protein (Luo et al., 2014;Liu et al., 2017;Courtney and Deiters, 2018).
This system relies on mutating the target protein at the desired codon to the infrequently used amber stop codon (UAG). The mutant gene is then transfected into cells along with plasmids encoding an engineered pyrrolysyl-tRNA synthetase/tRNA pair and the caged amino acid is added to the media. Here, a hydroxycoumarin-caged lysine (HCK) is transacylated onto the tRNA and thence ribosomally incorporated into the mutated gene in response to the UAG codon. In the context of LepB, the bulky hydroxycoumarin group incorporated onto lysine 39 blocks the active site (Dong et al., 2016). However, illumination of cells expressing this system with 405 nm light causes photolysis of the coumarin group, liberating the lysine residue, and generating wild-type, active LepB ( figure 2A). This provides precise and acute spatiotemporal control over LepB function in live human cells.
Optogenetic activation of LepB in cells expressing cPHx3 caused synthesis of PI(3,4)P2 on endosomes and striking recruitment of the cPHx3 probe ( figure 2B). This optogenetic activation was only observed in cells grown in the presence of exogenous HCK, and depended on cotransfection with the LepB K39UAG , ruling out off-target effects on endogenous UAG-containing genes or of light-exposure alone. Similarly, we observed clear translocation of PH-AKT to endosomes, which is expected given this PH domain's binding to both PIP3 and PI(3,4)P2 (Ebner et al., 2017). However, no recruitment of the PIP3-specific PH-ARNO 2G x2 was observed (Venkateswarlu et al., 1998;Manna et al., 2007), demonstrating that there was no further phosphorylation of the lipid at the 5-OH. Finally, we observed some depletion of the PI3P biosensor FYVE-EEA1 , though the biosensor remained mostly endosomebound, indicating a small fraction of total PI3P was converted to PI(3,4)P2 over the time course of these experiments (figure 2B).
Collectively, these results demonstrate that generation of PI(3,4)P2 in a cellular membrane is indeed sufficient to recruit cPHx3, which given its extensive in vitro selectivity (Dowler et al., 2000;Thomas et al., 2001), now passes all the key criteria that define a high-fidelity, genetically encoded biosensor for PI(3,4)P2 . The fidelity of the probe established, we next turned our attention to deploying this tool to answer some central questions about the lipid's metabolism and function.

Dominance of Class I PI3K in PI(3,4)P2 synthesis
The canonical view of PI(3,4)P2 synthesis induced by Class I PI3K signaling is that PI3K generates PIP3, which is then dephosphorylated at the 5-OH position by inositol polyphosphatase 5-phosphatase (INPP5) family members SHIP1 or SHIP2 (Vanhaesebroeck et al., 2012). Both enzymes are activated by tyrosine phosphorylation cascades initiated by the activated receptor tyrosine kinase. The extent of PI(3,4)P2 accumulation varies depending on receptor activation, with insulin or insulin-like growth factor reported to generate only modest amounts of the lipid Guilherme et al., 1996), which have not been detected with a single cPH probe (Kimber et al., 2002).
We tested the capacity of cPHx3 to detect insulin-induced PI(3,4)P2 production in HeLa cells ( Figure 3A). To unambiguously detect PIP3 in the same cells, we employed a tandem dimer of the PH domain from ARNO, specifically selecting the 2G splice variant with high PIP3 affinity (Cronin et al., 2004) and introducing the I303E mutation that prevents PH domain interaction with Arl GTPases (Hofmann et al., 2007). We selected this domain since it exhibits clear discrimination for PIP3 over PI(3,4)P2 (Manna et al., 2007). Insulin stimulation of HeLa cells generated a robust, transient increase in PIP3 biosensor at the PM, with a lagging, yet more sustained appearance of the PI(3,4)P2 biosensor (figure 3B, C) -a clear recapitulation of the original biochemical measurements of PI3K product generation (Stephens et al., 1991;Jackson et al., 1992;Hawkins et al., 1992), apparently supporting the canonical view.
As an alternative mechanism to activate Class I PI3K, we turned to chemically induced dimerization to recruit the inter-Src Homology 2 (iSH2) domain from the p85 regulatory subunit of PI3K (figure 3A). This system recruits endogenous PI3K p110 catalytic subunits to the membrane, inducing PIP3 synthesis (Suh et al., 2006). However, in this situation, tyrosine kinase-mediated activation of SHIP phosphatases is not expected. Nevertheless, iSH2 recruitment to the PM of COS-7 cells induced rapid PIP3 synthesis and the same, lagging PI(3,4)P2 accumulation (figure 3D). We therefore speculated that, in this context at least, PI(3,4)P2 accumulation might be driven by direct phosphorylation of PI4P by p110, given that the enzyme is known to perform this reaction in a test tube (Carpenter et al., 1990).
To test this speculation, we devised an experiment wherein the INPP5E enzyme would be corecruited to the PM in conjunction with iSH2. Since INPP5E will deplete both PI(4,5)P2 and PIP3, accumulation of PIP3 should be prevented ( figure 3E). PIP3 will be degraded into PI(3,4)P2 -but synthesis of PI(3,4)P2 will only be sustained if the p110 enzymes indeed directly converts PI4P into PI(3,4)P2. Compared with controls using a catalytically impaired INPP5E D556A mutant, INPP5E completely blocked the accumulation of the PIP3 biosensor at the PM after corecruitment (figure 3F). However, cPHx3 initially recruited to the PM at a similar rate to control, though its synthesis was rapidly cut off when INPP5E was co-recruited (figure 3G). Therefore, PI(3,4)P2 synthesis cannot be sustained in the absence of PIP3 generation, even under conditions where SHIP phosphatases are not predicted to become activated. Presumably, the PIP3 5-phosphatase activity responsible either comes from basal SHIP activity through interaction of its C2 domain with acidic PM lipids (Ong et al., 2007;Coq et al., 2017), or else from other INPP5 family members, which are all competent at converting PIP3 to PI(3,4)P2 (Trésaugues et al., 2014). Identification of the enzymes responsible will be a key question for future work.
Thus far, these experiments addressed the pool of PI(3,4)P2 generated downstream of Class I PI3K signaling. However, activity of class II PI3K have been shown to function in a more constitutive capacity (Posor et al., 2013;Marat et al., 2017). We therefore wanted to identify the source of PI(3,4)P2 that we observed at the PM in growing cells (figure 1). To this end, we employed the potent and Class I PI3K-selective inhibitor GDC-0941 (Kong et al., 2010) at 250 nM to distinguish Class I and Class II activities (figure 3H). After inhibitor treatment for 20 min, cells were subject to chemically-induced dimerization to recruit a co-expressed FKBP-INPP4B to the PM (figure 3I), the goal being to degrade any remaining PI(3,4)P2. As shown in figure 3I, GDC treatment led to the depletion of almost the entire pool of PM-associated PI(3,4)P2 from cells grown in serum. We cannot rule out that the small additional decrease induced in the 5 min after rapamycin addition is not an addition artifact ( figure 3I); area under the curve measurements for post rapamycin treatment vs. the 5 min period preceding it do not show a significant change (p = 0.43, Sidak's multiple comparisons test) as compared to the change following DMSO treatment (p < 10 -4 , Sidak's multiple comparison test after 2-way repeatedmeasures test, F = 61.9, p < 10 -4 ). However, the data do clearly indicate that the overwhelming majority of PI(3,4)P2 present in the PM of cells is generated by the class I PI3K pathway.

Direct Hydrolysis of PI(3,4)P2 by PTEN
So far, our data support a canonical view of Class I PI3K signaling, which is dominated by conversion of PI(4,5)P2 to PIP3, followed by degradation back to PI(4,5)P2 by PTEN or conversion to PI(3,4)P2 via INPP5 enzymes, most prominently SHIP1 and SHIP2 (Vanhaesebroeck et al., 2012;Fruman et al., 2017). PTEN and INPP5 thus represent a bifurcation of the pathway: in the former case, towards a simple inactivation; in the latter, the conversion to a modified but still active signaling state (Li and Marshall, 2015). PI(3,4)P2 is ultimately degraded to PI3P by INPP4A/B, a process intimately linked to endocytosis (Shin et al., 2005). However, it has been perplexing that the accumulation of PI(3,4)P2 seen after PI3K activation does not lead to a detectable increase in PI3P levels Stephens et al., 1991), implying an alternative route of degradation. Recently, Malek and colleagues have proposed that PTEN in fact directly converts PI(3,4)P2 to PI4P, terminating this signaling as well (Malek et al., 2017). The evidence was based on the requirement for PTEN knockout for EGFstimulated PI(3,4)P2 accumulation, in addition to showing that a PI(3,4)P2 3-phosphatase activity in MCF10 cell cytosol was lost in PTEN nulls. However, direct evidence for hydrolysis of PI(3,4)P2 in intact, living cells is still lacking.
The principle reason for the ongoing ambiguity over cellular activity of PTEN against PI(3,4)P2 has been the ambiguity when interpreting changes in lipid levels in cells. When PTEN loss induces PI(3,4)P2 accumulation, this can be explained by failure of PTEN to degrade PI(3,4)P2 directly -or alternatively, due to impaired PIP3 degradation, leaving more substrate available for the INPP5 enzymes (figure 4A). Careful mathematical modeling suggested the latter explanation did not explain the accumulation in MCF10 cells (Malek et al., 2017). Nonetheless, the evidence is indirect. We therefore devised an experiment utilizing the LepB PI3P 4-OH kinase. Since we showed that this enzyme generates an endosomal pool of PI ( signaling in addition to signals driven by PIP3 (Malek et al., 2017).
[text] We use this high fidelity and high sensitivity probe to develop the first direct evidence for the canonical pathway of PI(3,4)P2 synthesis by class I PI3K (i.e. by PIP3 de-phosphorylation, figure 3), first proposed nearly three decades ago (Stephens et al., 1991;Hawkins et al., 1992). We also provide direct evidence for a much more recent proposition (Malek et al., 2017): that the tumor suppressor PTEN also terminates PI(3,4)P2 signals in addition to those mediated by PIP3 (figure 4). We also generated the first optogenetically activatable lipid kinase, LepB (figure 2).
How PI(3,4)P2 signaling couples to different functions, and their relationship (or not) with the class I PI3K pathway, is still poorly understood (Li and Marshall, 2015;Hawkins and Stephens, 2016). This has largely been due to the difficulty in studying PI(3,4)P2 in isolation from the other, more abundant phosphoinositides such as PI(4,5)P2 and PIP3. However, recent advances in mass spectrometry are allowing sensitive detection of this lipid (Malek et al., 2017;Bui et al., 2018). We anticipate that our tools and cPHx3 probe will complement these approaches and greatly accelerate research in this area.
For now, the clearest demonstration is that most PI(3,4)P2 accumulating in cells is derived from INPP5 activity on class I PI3K-synthesized PIP3. Without stimulation, resting levels of PI(3,4)P2 are extremely low (Stephens et al., 1991;Hawkins et al., 1992), with immunocytochemical estimates suggesting that perhaps 40% of this is generated by the class II PI3K-C2α . The function of the greatly expanded class I PI3K-driven pool remains an ongoing question. Conversion of PIP3 to PI(3,4)P2 will terminate signals generated by PIP3-selective effectors, maintain signals by more promiscuous effectors (like Akt), and initiate signaling by PI(3,4)P2-selective effectors (Hawkins and Stephens, 2016). What is the purpose of this tiered lipid signaling system? A clue comes from perhaps the best characterized PI(3,4)P2-selective effector protein: the TAPP proteins from which our probe is derived. Mice harboring point mutations in Plekha1 and Plekha2 that disrupt PI(3,4)P2-binding of the TAPP1 and TAPP2 cPH domains exhibit augmented PI3K signaling through insulin and B-cell receptors (Gray and Alessi, 2011;Landego et al., 2012), suggesting a major function of the lipid is in negative feedback of class I PI3K signaling. Interestingly, PI(3,4)P2 is also implicated in endocytosis (Shin et al., 2005;Posor et al., 2013;Boucrot et al., 2015). Therefore, it could be possible that PI(3,4)P2 downregulates PI3K signals by triggering internalization of activated receptors.
Such a model may reconcile conflicting data over the generation of PI(3,4)P2 in clathrin mediated endocytosis. The class II PI3K-C2α was shown to generate a pool of PI(3,4)P2 essential for constitutive endocytosis (Posor et al., 2013), whereas a co-incidence detector probe showed that the enzyme was dispensable for PI(3,4)P2 accumulation at clathrin-coated structures (He et al., 2017). Although seemingly at odds, it is worth noting that careful quantitative modeling of lipid dynamics during formation of the clathrin bud found that just enough PI(3,4)P2 is produced to recruit its SNX9 effector protein -meaning that the level of free lipid might be extremely low and invisible to even the most sensitive biosensor (Schöneberg et al., 2017). So why was PI(3,4)P2 observed at clathrin structures at all (He et al., 2017)? We propose that the greatly expanded pool of class I PI3K synthesized PI(3,4)P2 was being detected. Indeed, the cells in He et al were imaged in 5% serum, and most experiments were conducted in SUM159 cells, which have constitutive class I PI3K activation via the oncogenic 1047A mutation in p110α (Barnabas and Cohen, 2013). Intriguingly, PI(3,4)P2 synthesis by class I PI3K activity was also found to be essential for clathrin-independent, fast endophilin mediated endocytosis (Boucrot et al., 2015). An attractive model is therefore that PI3K-C2α produces a minor pool of PI(3,4)P2 that supports constitutive clathrin-mediated endocytosis, whereas greatly expanded pools downstream of Class I PI3K are required for accelerated clathrin-mediated and independent endocytosis after activation, whilst also sustaining Akt signaling.
A second major finding is that PI(3,4)P2 is directly hydrolyzed by PTEN (Malek et al., 2017). This result is surprising, since meticulous work with purified protein found much poorer PTEN activity against PI(3,4)P2 vs. PIP3 (McConnachie et al., 2003). The discrepancy likely derives from the very different enzyme-substrate interactions in living cells; indeed, turnover numbers for PTEN in living cells are several orders of magnitude higher than in the test tube (McConnachie et al., 2003;Feng et al., 2014). It therefore seems likely that PTEN regulates PI(3,4)P2 signaling in addition to PIP3. Intriguingly, the recent observations that PTEN reduced signaling from forming clathrin-coated structures and slows their maturation is entirely consistent with the roles of both class I and class II PI3K proposed in the previous paragraph (Rosselli-Murai et al., 2018).
Finally, since PTEN's activity on PI(3,4)P2 is the direct reversal of the PI4P conversion mediated by class II PI3K, and since PTEN has recently been found to localize to endosomal structures (Naguib et al., 2015), PTEN seems poised to regulate PI3K-C2β signaling from late endosomes/lysosomes too (Marat et al., 2017).
In summary, we have developed and validated a sensitive genetically encoded lipid biosensor for PI(3,4)P2. We show this lipid accumulates in cells primarily in response to class I PI3K activity, via circuitous dephosphorylation of PIP3. We also provide evidence for direct hydrolysis of PI(3,4)P2 by PTEN. Collectively, these results provide fresh impetus to dissect the physiological and pathological signaling outcomes driven by PI(3,4)P2, and provides a powerful new tool to aid in this endeavor.

Materials and Methods
Cell Culture and transfection COS-7 (ATCC CRL-1651)  Plasmids pPAmCherry-C1 (Addgene plasmid 31929) was a kind gift of Vladislav Verkhusha (Subach et al., 2009). EGFP (Aequorea victoria GFP with F64L and S65T mutations with human codon optimization), mCherry, Rhodopseudomonas palustris bacteriophytochrome BphP2-derived near-infrared fluorescent protein variant iRFP713 and Entacmaea quadricolor GFP-like protein eqFP578 variant pTagBFP2 were used in the Clontech pEGFP-C1 and N1 backbones as described previously (Zewe et al., 2018). TAPP1 cPH and LepB sequences were obtained as synthetic gblocks (IDT). All plasmids were verified by dideoxy sequencing. Constructs generated in this study are freely distributed through Addgene. Plasmids were constructed using standard restriction cloning or NEB HiFi assembly (New England Biolabs E5520S), or else obtained from the sources indicated below:  (Luo et al., 2014) were stored at -20˚C in DMSO at 100 mM.

Image Analysis
Images in Nikon nd2 image format were imported into the open access image analysis package Fiji (Schindelin et al.), using the LOCI BioFormats importer (Linkert et al., 2010). A custom written macro was used to combine fields into a single enlarged image for the purposes of image analysis (though never for presentation).

Quantifying the difference in intensities between EGFP and iRFP channels (figure 1B&C):
Regions of interest (ROI) were drawn around each cell, and a custom-written macro was used to measure the average pixel intensity in these ROI and then normalize each pixel to this value.
The normalized iRFP (cytosolic) image was then subtracted from the normalized EGFP-cPH channel to yield the subtracted image.

Intensity changes in specific compartments imaged by confocal (figures 2B, 4B & C)
: Cells were measured inside ROI. A second image channel (mCherry-Rab5 in figure 2 and LAMP1-FRB-iRFP in figure 4) was auto-thresholded and used to generate a mask to measure the normalized pixel intensity of the EGFP channel, as previously described in detail (Zewe et al., 2018). tracked using the open-source Fiji implementation of the "u-track" single particle tracking algorithm (Jaqaman et al., 2008). A "Difference of Gaussians" filter was used with a hard threshold and an estimated dimeter of 0.5 µm for the estimated diameter (i.e. an approx 8x8 pixel neighborhood) of single molecules. Co-ordinates of the trajectories were exported as xml files, and mean-square displacement at different time lags (Vrljic et al., 2007) was calculated along with trajectory lifetime distributions using custom written code in Python.
Selecting representative images for presentation: example images were selected based on having the best signal to noise possible, whilst also having measured values close to the sampled population median, and are always within the central interquartile range.
Data presentation and statistics was performed using Graphpad Prism 7. Data were subject to the D'Agostino and Pearson normality test to select for parametric or non-parametric tests.