Nir1-LNS2 is a novel phosphatidic acid biosensor that reveals mechanisms of lipid production

  1. Claire C. Weckerly
  2. Taylor A. Rahn
  3. Max Ehrlich
  4. Rachel C. Wills
  5. Joshua G. Pemberton
  6. Michael V. Airola
  7. Gerald R. V. Hammond

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Abstract

Despite various roles of phosphatidic acid (PA) in cellular functions such as lipid homeostasis and vesicular trafficking, there is a lack of high-affinity tools to study PA in live cells. After analysis of the predicted structure of the LNS2 domain in the lipid transfer protein Nir1, we suspected that this domain could serve as a novel PA biosensor. We created a fluorescently tagged Nir1-LNS2 construct and then performed liposome binding assays as well as pharmacological and genetic manipulations of HEK293A cells to determine how specific lipids affect the interaction of Nir1-LNS2 with membranes. We found that Nir1-LNS2 bound to both PA and PIP 2 in vitro . Interestingly, only PA was necessary and sufficient to localize Nir1-LNS2 to membranes in cells. Nir1-LNS2 also showed a heightened responsiveness to PA when compared to biosensors using the Spo20 PA binding domain (PABD). Nir1-LNS2’s high sensitivity revealed a modest but discernible contribution of PLD to PA production downstream of muscarinic receptors, which has not been visualized with previous Spo20-based probes. In summary, Nir1-LNS2 emerges as a versatile and sensitive biosensor, offering researchers a new powerful tool for real-time investigation of PA dynamics in live cells.

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    Manuscript number: RC-2024-02413R

    Corresponding author(s): Hammond, Gerald

    1. General Statements [optional]

    *We are grateful to the three reviewers for such thorough and thoughtful comments. Data or re-writes that we have on hand that address many of these comments have been incorporated already. We also have a comprehensive experimental plan to address all of the remaining major comments. Reviewer’s comments are in light italics, whilst our responses appear in regular font below. We added reviewer numbering for ease of cross-reference to the original comments, with the format: reviewer X’s comment number N as *#X.N

    *Overall, we were thrilled that the reviewers agreed that our work is of significance and broad interest: *

    “The development of a new, superior PA-sensor is a significant advance in the fields of lipid signaling and specific lipid-protein interactions, that will benefit research on lipid-mediated cellular signaling and intracellular lipid trafficking.” – reviewer 1.

    “The lipid biology community would be highly interested in using the new PA-binding tool to study lipid localization in live cells.” – reviewer 2.

    “Tracking intracellular phosphatidic acid (PA) in live cells is essential for understanding its cellular functions, leading to the development of genetically encoded lipid biosensors. While several PA biosensors have been developed, they often suffer from limited sensitivity or specificity.” – reviewer 3

    2. Description of the planned revisions

    #1.2: P 4 and Fig. 2 mention a 'novel domain structure', responsible for binding of PA and PIP2 (in vitro). What exactly is this novel domain structure? Why have the two parts of Nir1-LNS, the AAH and the 263 amino acid domain, not been tested in similar liposome assays as in Fig. 2A? Lipid binding in vivo, as tested in the experiment of Fig. 2D, is confounded by endogenous PA binding proteins.PA is expected to bind the SIDGS motif, as this is conserved from the Lipin catalytic motif (p. 5). However, experimental evidence of this appears to be lacking. Nevertheless, it is several times presented as a fact in the text. Pag. 7: 'This suggests that the SIDGS motif alone is not the sole PA binding pocket as the LNS2 domain requires both that motif and the amphipathic helix for sustained binding to membrane embedded PA.' Pag. 13: '... One reason for this difference could be that the Nir1-LNS2 is not a novel bona fide PA binding pocket. Rather, it requires both the SIDGS motif and an N-terminal amphipathic helix for membrane interactions (Figure 2). These conclusions therefore appear to be not accurate and would need te be rephrased. If the authors could show that PA binding by Nir1-LNS can be eliminated by mutating residues in the SIDGS motif, this would not only substantiate the above claims, but also make for a negative control protein, next to Nir1-LNS2. For future applications of Nir1-LNS2 as PA biosensor in other organisms this would be useful.

    In addition to the already included data on cellular binding of the R784E mutant, we do plan to test this variant in the liposome binding assays to show loss of PA binding abilities as the reviewer has suggested. We also plan to evaluate proper folding of the R784 mutant through circular dichroism.

    #1.7. Fig. 2A suggests cooperativity in binding of Nir1-LNS2 to PA-containing liposomes. Please mention/comment! Does binding to PIP2-containing liposomes also exhibit cooperativity?

    Using a nonlinear fit, we were able to determine Hill coefficients for PA binding. This has now been included in Figure 2B (formerly Figure 2A) and the following text has been added to page 6, left column, third paragraph: “In addition, we found that Nir1-LNS2 bound PA-rich liposomes in a concentration dependent manner (Figure 2B). By fitting the binding curve to this data, we found that the interaction of Nir1-LNS2 with PA provided a Kd value of ~19 mol%. Interestingly, Nir1-LNS2 binds to PA in a highly cooperative manner. The Hill coefficient for the interaction of Nir1-LNS2 with PA was calculated to be approximately 4 (Figure 2B).”

    However, due to the liposome binding assay used that utilizes a set total lipid concentration but alters mol% lipids, the Kd that we determined is not a “traditional” Kd. Therefore, we plan to repeat this assay using constant PA concentrations but increasing total concentrations of lipid so that we can make a better fit and get a more accurate Kd value and Hill coefficient. We also plan to do the same assay with PIP2 to determine Kd values and Hill coefficients for that interaction.

    #2.2. The authors mention high affinity of Nir1-LNS2, but it lacks in vitro characterization that should demonstrate the higher affinity of Nir1-LNS2 compared to conventional probes such as Spo20. The authors should perform side-by-side comparison in Fig 2 to compare the PA affinity and specificity of Nir1-LNS2 compared to Spo20.

    We plan to take the reviewer’s advice and directly compare Spo20-PABDx2 (and/or the single PABD depending on what we can get to purify correctly) and Nir1-LNS2 in the liposome binding assay.

    Additionally, we propose to further characterize these sensors in cells as well. To start, we have added a direct comparison of the Spo20-PABDx2 and Nir1-LNS2 response to PA production at the PM (by PMA stimulation) and at mitochondria (by FKBP-DGKa) in Figure 4. The text has been updated to reflect this on p.10, left paragraph, 3rd paragraph: “Importantly, we also observed that Nir1-LNS2 responds to this ectopic PA production quicker and more robustly than NES-PABDx2-Spo20 does, as can be seen when the responses from Figure 4F are plotted together (Figure 4H). When analyzing the responses to PA production at the PM by PMA stimulation in Figure 1D and Figure 1F, we similarly see that the Nir1-LNS2 translocates to the PM more robustly and in a shorter timeframe (Figure 4G). This suggests that the Nir1-LNS2 can serve as a high affinity PA biosensor at various cellular locations.”

    Furthermore, as suggested in the “General Assessment” we propose to use the FKBP-DGKa system to produce PA on other organelles such as the Golgi and ER and then we can directly compare the response of Spo20-PABDx2 and Nir1-LNS2 to the increase in PA at these organelles. This data will be added to Figure 4 for a full comparison of the sensors across cellular locations.

    #2.4. Fig 3 and Fig 4 need the validation of PIP depletion/production using PIP-binding probes.

    We propose to repeat the experiments using TIRF in figure 3 as it will give us increases sensitivity, and also compare selectivity with the currently used spo20-based biosensors.

    #2.“General assessment”: The existing PA-binding probe using Spo20 is indeed quite blunt, which takes minutes to see appreciable accumulation of the probe upon PA production. Nir1-LNS2 can be indeed useful if it offers better spatiotemporal precision. However, the advantage of this tool over existing tools is not convincing without head-to-head comparison of either (1) in vitro characterization of PA binding affinity and selectivity between Nir1-LNS2 and Spo20 or (2) response to PA produced on different subcellular localizations other than plasma membrane and mitochondria (e.g., endosomes, golgi, and endoplasmic reticulum).

    In order to address the selectivity of Nir1-LNS2 and Spo20, we propose to repeat the experiments in Figure 3 with the PJ enzymes in order to see how the PM PIPs affect Spo20 membrane binding, as described in our response to #2.4. Previously published data, as well as our own unpublished observations suggest that Spo20 interacts with the anionic PIPs to a greater extent than Nir1-LNS2 does (Nakanishi et al., 2004 doi: 10.1091/mbc.e03-11-0798; Horchani et al., 2014 doi: 10.1371/journal.pone.0113484). If we can show that Spo20’s interactions with the PM are significantly influenced by the PIPs, then this will add more evidence to the idea that Nir1-LNS2 is more selective for PA.

    As described in response to #2.2, we are also planning a side-by-side comparison of spo20 based protein binding on liposomes alongside Nir1-LNS2.

    Also, as discussed above, we agree with the reviewer that looking at the Nir1-LNS2 and Spo20 responses to PA production at other organelles would increase confidence that Nir1-LNS2 has a higher affinity for PA. We propose to add these experiments to Figure 4.

    #3.1. The direct measurement of the binding affinity of Nir1-LNS2 with PA, e.g., Kd, is essential; this information will help the field explore the potential usage of Nir1-LNS2.

    Using a nonlinear fit, we were able to determine Hill coefficients for PA binding. This has now been included in Figure 2B (formerly Figure 2A) and the following text has been added to p.6, left column, 3rd paragraph: “In addition, we found that Nir1-LNS2 bound PA-rich liposomes in a concentration dependent manner (Figure 2B). By fitting a nonlinear curve to this data, we found that the interaction of Nir1-LNS2 with PA provided a Kd value of ~19 mol%. Interestingly, Nir1-LNS2 binds to PA in a highly cooperative manner. The Hill coefficient for the interaction of Nir1-LNS2 with PA was calculated to be 4.323 (Figure 2B)…. This suggests that the amphipathic helix and the SIDGS-containing domain may both interact with the membrane leading to the cooperative nature of Nir1-LNS2’s binding of PA-rich liposomes (Figure 2B).”

    However, due to the liposome binding assay used that utilizes a set total lipid concentration but alters mol% lipids, the Kd that we determined is not a “traditional” Kd. Therefore, we plan to repeat this assay using higher total lipid concentrations with a fixed PA mol% so that we can make a better fit and get a more accurate Kd value and Hill coefficient. Furthermore, we plan to directly compare Spo20-PABDx2 (and/or the single PABD depending on what we can get to purify correctly) and Nir1-LNS2 in the liposome binding assay to directly compare their affinities for PA in vitro, as described in response to #2.2.

    #3.2. As mentioned by the authors, there is a confusing inconsistency regarding why Nir1-LNS2 binds to PIP2 in vitro but not in cells. Going beyond what has been discussed in the manuscript, there is a possibility that PIP2 could induce Nir1-LNS2 aggregation, leading to pelleting after centrifugation, among many other possibilities. I recommend the authors perform additional in vitro experiments, including but not limited to the liposome floatation assay to directly examine Nir1-LNS2 binding to the liposomes with varied compositions.

    This is an excellent suggestion. We plan to check for aggregation by liposome flotation with Nir1-LNS2 in the presence of high mol% of PA and PIP2. In addition, we will also perform circular dichroism to see if PA or PIP2 liposomes are inducing any unfolding of Nir1-LNS2.

    #3.3. In Fig. 2D, it would be beneficial to examine the constructs Nir1-613-630 and Nir1-631-894, comparing them with Nir1-LNS2 using liposome sedimentation and floatation assays to evaluate the contribution of the SIDGS motif and the amphipathic helix in binding PA.

    Per our response to #1.2, we looked around the SIDGS motif to find the residue that would mediate the binding of membrane embedded PA, which our data suggests is R784 (Figure 2D). We do plan to test the R784 mutant in the liposome binding assays to show loss of PA binding abilities as the reviewer has suggested. We also plan to evaluate proper folding of the R784 mutant through circular dichroism.

    3. Description of the revisions that have already been incorporated in the transferred manuscript

    #1.1: CCH treatment of HEK293A cells leads to the PM localization of the DAG sensor C1ab-Prkd1 as well as Nir1-LNS2 (Fig. 5), and the kinetics of these changes - Nir1-LNS2 would lag behind C1ab-Prkd1 fluorescence - is taken as evidence for Nir1-LNS2's specific binding of PA rather than DAG: Pag. 10: 'When we look at the first 2-minutes after CCh addition, we see that C1ab-Prkd1 moves to the PM much faster than Nir1-LNS2 does (Figure 5D). The delay in Nir1-LNS2 translocation makes sense given DAG is produced first and then converted into PA, again indicating that Nir1-LNS2 is specific for PA. 'Fig. 5 legend: 'The Nir1-LNS2 response to PLC depends on PA and not DAG.(...) (D) Nir1-LNS2 translocation to the PM (data replicated from Figure 5C) lags behind C1ab-Prkd1 (data replicated from Figure 5B) in response to CCh addition. 'The validity of the conclusion from this experiment seems questionable. The argument relies on the low values of Nir1-LNS2's normalized fluorescence intensity compared to C1ab-Prkd1's, in the first two minutes of stimulation, when DAG is expected to accumulate (Fig. 5D). However, if Nir1-LNS2 would bind DAG, the resulting fluorescence is expected to be low, i.e. compared to the much higher signal resulting from subsequent PA binding. Moreover, in this system, which co-expresses the two sensor proteins, competition in binding may account for the apparent precedence of one sensor over the other. Thus, even if the increase in C1ab-Prkd1 fluorescence would precede Nir1-LNS2 - following the authors' interpretation - this would not exclude binding of DAG by Nir1-LNS2.Fig. 5D shows the confocal images of cells after 30 sec CCH treatment using the two sensors, next to the respective controls, replicated from Fig. 5B/C. However, in Fig. 5D, different colors are used for Nir1-LNS2 than in Fig. 5C, which makes comparison along the time course difficult. In conclusion, the data presented in Fig. 5 do not exclude DAG binding by Nir1-LNS2, and modification of the conclusions from this experiment throughout the ms (including the cited sentences) is recommended. Consider removal of Figure 5D. Despite these considerations, the authors' final conclusion regarding the specificity of Nir1-LNS2 towards PA appears well supported (e.g. by the data presented in Figure 4).

    We agree with the reviewer that the interpretation of the kinetic data is ambiguous and does not fully negate the idea that Nir1-LNS2 may bind to DAG. We have modified the interpretation accordingly. However, we have left the kinetic comparison of the DAG vs Nir1-LNS2 biosensors since these reflect the expected dynamics of the two lipids downstream of PLC. The data are now interpreted as follows on p. 10, right column, second paragraph: “The PM accumulation lagged that of DAG, consistent with conversion of DAG to PA by DGKs (Figure. 5D). Alternatively, in cells treated with CCh and then atropine, Nir1-LNS2 localized to the PM after CCh was added but was then observed returning to the cytoplasm over the 15-minute treatment with atropine as PA levels declined (Figure 5C). Overall, this experiment shows that Nir1-LNS2 binding to the PM follows the expected kinetic profile of DGK-produced PA.” Likewise, the legend for figure 5 is now labelled “Nir1-LNS2 detects PLC stimulated PA production.” to remove explicit conclusions about PA vs DAG binding.

    #1.2: P 4 and Fig. 2 mention a 'novel domain structure', responsible for binding of PA and PIP2 (in vitro). What exactly is this novel domain structure? Why have the two parts of Nir1-LNS, the AAH and the 263 amino acid domain, not been tested in similar liposome assays as in Fig. 2A? Lipid binding in vivo, as tested in the experiment of Fig. 2D, is confounded by endogenous PA binding proteins.PA is expected to bind the SIDGS motif, as this is conserved from the Lipin catalytic motif (p. 5). However, experimental evidence of this appears to be lacking. Nevertheless, it is several times presented as a fact in the text. Pag. 7: 'This suggests that the SIDGS motif alone is not the sole PA binding pocket as the LNS2 domain requires both that motif and the amphipathic helix for sustained binding to membrane embedded PA.' Pag. 13: '... One reason for this difference could be that the Nir1-LNS2 is not a novel bona fide PA binding pocket. Rather, it requires both the SIDGS motif and an N-terminal amphipathic helix for membrane interactions (Figure 2). These conclusions therefore appear to be not accurate and would need te be rephrased. If the authors could show that PA binding by Nir1-LNS can be eliminated by mutating residues in the SIDGS motif, this would not only substantiate the above claims, but also make for a negative control protein, next to Nir1-LNS2. For future applications of Nir1-LNS2 as PA biosensor in other organisms this would be useful.

    We have clarified that the domain architecture of the Nir1-LNS2 is not a novel domain structure generally, but novel for a PA binding protein which are typically just helices such as that seen in Spo20. Figure 2 is now titled “Nir1-LNS2 shows specificity for PA and PIP2 in vitro, based on a novel PA-binding domain.”

    We have also clarified that the SIDGS motif is not the actual location of PA binding, but rather is only the motif conserved with the Lipin/Pah active site. R784 appears to be a PA coordinating residue near the SIDGS, as the positive residue can interact with the negative lipid. Furthermore, we agreed with the reviewer that mutating this residue to perturb PA binding was a much more convincing experiment. We have now included this data in Figure 2 and rewritten the following passages.

    From Page 6, left column, last paragraph: “The putative Lipin catalytic motif DxDxT is partially conserved in Nir1-LNS2 as a SIDGS motif spanning residues 742-746. We looked for positively charged residues nearby that could bind to the PA in the membrane and coordinate its entrance into the SIDGS site. The active site of the Lipins has a nearby Arg residue which was predicted to perform this role (Khayyo et al., 2020). AlphaFold analysis of Nir1-LNS2 showed that this residue was also conserved in Nir1-LNS2 as R784, and that the side chain of the Arg sticks out toward the membrane interface where it would be able to contact the negatively charged PA (Figure 2C).

    The conservation of these features between the Lipins and Nir1-LNS2 suggests that PA binds this positively charged residue near the SIDGS pocket within Nir1-LNS2 (Kim et al., 2013; Khayyo et al., 2020). However, for efficient catalytic activity, the Lipins also require an N-terminal amphipathic helix for membrane interaction. This helix is made up of residues 1-18 in Tetrahymena thermophila Pah2 (Khayyo et al., 2020), and residues 613-630 in the N-terminus of Nir1-LNS2 are predicted to form a similar amphipathic helix (Figure 2C). We therefore tested whether the N-terminal helix of Nir1-LNS2 was necessary for interaction with PA at the PM. We made two truncations of the Nir1-LNS2 construct: Nir1-613-630 is the isolated amphipathic helix, while Nir1-631-894 is the rest of the domain excluding the helix but including the SIDGS motif. Surprisingly, neither truncated construct responded to PMA by binding the PM, and they even showed reduced basal PM localization (Figure 2D).

    Although Figure 2D suggests that the SIDGS motif alone is not sufficient for membrane interactions, we probed into the suspected PA binding residue R784 by mutating it into a negatively charged Glu residue, which should disrupt its interaction with the negatively charged lipid. The R784E mutation completely ablated Nir1-LNS2 interactions at the PM after PMA stimulation and showed reduced association with the PM even before PMA stimulation (Figure 2D).

    Altogether, our data suggests that the LNS2 domain requires both the larger SIDGS-containing domain and the amphipathic helix for sustained binding to membrane-embedded PA, but that the PA may directly interact with R784 near the SIDGS motif. Therefore, the Nir1-LNS2 provides a novel PA binding domain with a tertiary structure beyond the simple amphipathic helices found in Spo20.”

    We have also rewritten this sentence in the discussion, p. 14, right column, second paragraph: “As far as the use of Nir1-LNS2 as a biosensor, the one caveat is the discrepancy in its specificity: in vitro PA and PIP2 were sufficient to recruit Nir1-LNS2 to PC liposomes (Figure 2), but in vivo only PA was sufficient for mitochondrial recruitment (Figure 4). One reason for this difference could be that the Nir1-LNS2 requires R784 near the SIDGS pocket and an N-terminal amphipathic helix for membrane interactions (Figure 2).”

    #1.3. Pag. 7: '...small caveat to consider when using Nir1-LNS2 to study PA, the data also demonstrates that Nir1-LNS2 is not specifically interacting with any of the PM PIPs in cellular membranes.'This seems not accurate, since the data in Fig. 3 suggest that PI4P could be involved in membrane localization of Nir1-LNS2. It remains however unresolved whether this is a specific interaction with this PIP.

    We have rewritten the text on Page 8, right column, first paragraph accordingly: “This data suggests that decreasing the anionic charge of the membrane through depletion of PIPs slightly reduces Nir1-LNS2’s ability to interact with the PM, but it doesn’t fully re-localize the sensor. Therefore, this is a caveat to consider when using Nir1-LNS2 to study PA.”

    #1.4. Please note that the presence of PE increases the ionization (and negative charge) of PIP2 (Graber et al., 2012) rather than dilutes the negative charge as stated in the Discussion on p.13. Please revise!

    We have updated the text on Page 14, right column, last paragraph: “The presence of other lipids such as PI, the formation of PIP2-rich domains, and even interactions with neighboring proteins can increase hydrogen bonding of PIP2and dilute the negative charge (Graber et al., 2012; Borges-Araújo and Fernandes, 2020). Phosphatidylethanolamine on the other hand, increases PIP2 ionization and its negative charge, though these effects are also thought to be reduced by PIP2 intramolecular hydrogen bonding which competes for the charges on the lipid (Graber et al., 2012).”

    #1.5. P 1 The authors may consider adding a 4th criterium for a lipid biosensor: the sensor should not serve as a sink for the lipid by removing/sequestering it from the active pool, thereby interfering with other interactions/conversions.

    We agree that biosensors should not sequester a significant fraction of their cognate lipids and affect downstream pathways by competing with endogenous binding partners. We have rewritten the following text regarding Figure 6 to make this distinction more clear:

    Page 13, left column, last paragraph: “As Nir1-LNS2 shows high affinity for PA across cell lines, this brings up the concern that use of Nir1-LNS2 will sequester PA and inhibit endogenous signaling pathways that depend on PA…Therefore, we conclude that use of Nir1-LNS2 as a PA biosensor does not sequester significant amounts of PA. It is suggested that cellular homeostasis may compensate for the amount of bound lipid by increasing synthesis of free lipid, as this has been seen with the PIP2 biosensor PH-PLCd1 (Traynor-Kaplan et al., 2017). While PA has a plethora of cellular functions, the fact that Nir1-LNS2 expression does not disrupt MCS formation shows promise that the high affinity of Nir1-LNS2 will not inhibit downstream PA signaling.

    #1.6. Nir1 lacks a PITP domain (Fig. 1), yet is referred to as lipid transfer protein: please elaborate/explain.

    The following text has been added to Page 2, left column, last paragraph to clarify this point: “This family of proteins, made up of Nir1, Nir2, and Nir3, form ER-PM membrane contact sites (MCS) to exchange PA and phosphatidylinositol (PI) between the compartments (Cockcroft and Raghu, 2016; Kim et al., 2015). While Nir1 lacks a functional PITP domain, it was initially classified as part of the PITP family based on the homology of its other domains with Nir2 and Nir3. Furthermore, Nir1 has a role in lipid transfer by facilitating Nir2 recruitment to the MCS (Quintanilla et al., 2022).”

    #1.8. Indicate the concentrations of PC and protein in the legend to Fig. 2 panels A and B. M&M says 2 mM PC, according to the PA-concentrations above panel 2A, this should be 1 mM. Please clarify.

    We have corrected the typo in panel 2A (now panel 2B) and have updated the Figure 2 legend as follows, “(A) A representative SDS-PAGE gel is shown for Nir1-LNS2 binding to various PM lipids in POPC liposomes. (B) A representative SDS-PAGE gel is shown for Nir1-LNS2 binding of increasing PA molar concentrations in POPC liposomes. For both A and B, the lipids indicated were mixed with POPC to produce a 2 mM solution, then 50 uL of the resulting liposome mixture was incubated with 50 uL of Nir1-LNS2 at ~1 mg/mL. Supernatant (S) and pellet (P) lanes were quantified using ImageJ to determine percent protein bound. The protein-only control pellet was used as a baseline (input). Nir1-LNS2 appears on the gel at 37 kDa.”

    #1.9. In Fig. 2B, PI is missing. Any specific reason?

    We have updated the text on Page 6, left column, second paragraph to discuss the low levels of PI at the PM, which is why we did not include this lipid. “Using this same PC background, we tested the efficacy of the PM lipids DAG, PA, PS, PI4P and PIP2 in recruiting Nir1-LNS2 to membranes. While PI serves as a substrate for PI4P and PIP2 synthesis (collectively referred to as the phosphatidylinositol phosphates (PIPs)) at the PM, levels of PI at the PM are very low compared to the PIPs and therefore PI itself was not tested (Zewe et al., 2020; Pemberton et al., 2020).”

    #1.10. Move Fig. 2C to the Introduction and extend it to illustrate the shared conserved features of Nir1-LNS2 and lipin.

    We would like to keep the diagram of the Nir1-LNS2 in Figure 2 where the features are discussed in more detail than in the introduction. However, we did add this sentence to the introduction __on p.2, right column, second paragraph __that refers the reader to the cartoon in Figure 2. “These features are conserved in the Nir LNS2 domains, except for the catalytic Asp in the DxDxT motif and another Mg2+-coordinating residue (Figure 2C).”

    #1.11. P 13. 'While real-time IMPACT does not directly report on PA levels as it does not use the endogenous PLD substrate PC, ...'It is true that this method doesn't directly report on PA levels, but that is because it uses a click chemistry probe as substrate for PLD's transphosphatidylation reaction. Contrary though to what is stated by the authors, this reaction still uses the endogenous substrate of PLD, PC (Liang et al. 2019; www.pnas.org/cgi/doi/10.1073/pnas.1903949116).

    We have rewritten the aforementioned sentence in the discussion (Page 15, right column, second paragraph): “While real-time IMPACT does not directly report on PA levels as it creates a unique fluorescent lipid, it offers several advantages such as being able to interrogate lipid trafficking over time.”

    #1.12. Fig. 1K: Control must have been treated with PMA plus vehicle (DMSO); if so, please indicate that vehicle was added.

    Figure 1K and its legend have been updated. The legend now reads “Stimulating HEK293A cells with 100 nM PMA and 750 nM of the PLD inhibitor FIPI reduces the Nir1-LNS2 response to PMA and cell media (Veh)”

    #1.13. Figure 6C: How is DFt/Fpre defined? Add to legend.

    We have updated the Figure 6 legend to read “MCS formation was quantified as the change in fluorescence at a given time (Ft) divided by the fluorescence before CCh stimulation (Fpre).”

    #1.14. P 4: 'The boundaries of the Nir2-LNS2 (Uniprot: O00562) and Nir3-LNS2 (Uniprot: Q9BZ72) were also defined using AlphaFold predictions of the structure of these domains.' How was the extent of each of these domains determined - they are much larger than the previously published sequences of LNS2 domains (Kim et al. 2013; Embo Rep. 14:891-899. doi:10.1038/embor.2013.113)?

    We have clarified the definition of boundaries by updating the following sentence on Page 4, right column, 5thparagraph: “The boundaries of the Nir2-LNS2 (Uniprot: O00562) and Nir3-LNS2 (Uniprot: Q9BZ72) were also defined using AlphaFold predictions of the structure of these domains. Previous definitions of the Nir2-LNS2 domain have considered the domain smaller than we do here (Kim et al., 2013, 2015) . However, according to AlphaFold, the boundaries set previously exclude a large N-terminal beta barrel that is conserved in the Lipin/Pah PAPs, as well as disrupt the domain fold that is homologous to the Lipin active site. Therefore, we are confident that our constructs include the entire LNS2 fold.”

    #1.15. In Fig. 3 legend, specify the starting condition of Nir1-LNS2 binding?? Which fluorescence are we looking at?

    This figure has now been revised in response to __point #3.5, __which hopefully also clarified this point.

    #1.16. In legend to Fig. 6 please specify the fluorescent tags used. Have they been shown not to affect protein function?

    We have updated the figure legend to specify that GFP-Nir2 was used in conjunction with iRFP-Nir1-LNS2, we also changed the text on Page 13, right column, second paragraph that refers to this experiment. It now reads “We co-expressed a GFP-tagged Nir2 and either iRFP-Nir1-LNS2 or iRFP-TubbyC, a PIP2 biosensor that is not expected to affect MCS formation. It should be noted that although we have used the NG-tagged Nir1-LNS2 the most extensively, the iRFP and mCherry-tagged biosensors have behaved the same as the NG-tagged version in the experiments where we utilized them.”

    #2.1. Nir1-LNS2 seems to show variable basal localization across different representative images presented in the manuscript. A part of them were justified by the effect of other anionic species by PIP (such as Fig 3 where they co-expressed PIP-degrading enzymes). For example, cells in Fig 1F and those in Fig 4F show quite different basal localization of Nir1-LNS2. Is it due to difference in expression level, cell conditions, or other factors The significant amount of plasma membrane basal localization seems to indicate that Nir1-LNS2 localization is affected by its binding to PI(4,5)P2.? The significant and potentially variable plasma membrane localization of Nir1-LNS1 can limit the utility of this probe.

    We have added __Supplemental Figure 1 __to show the range of Nir1-LNS2 basal localization compared to NES-PABDx2-Spo20 and PASS. We believe that this localization is due to variable amounts of basal PA combined with some non-selective anionic interactions at the PM. The following paragraph has been added to page 7, left column, first paragraph to discuss this point, “Since the R784E mutant showed reduced basal PM localization, we wanted to further characterize the basal localization of the wild-type Nir1-LNS2. The basal localization of wild-type Nir1-LNS2 varies somewhat between cells, but analysis of all of the cells used throughout this study determines that the basal PM/Cyt ratio of the wild-type Nir1-LNS2 is 1.0644 ± 0.0672, which suggests that at resting conditions Nir1-LNS2 is slightly enriched at the PM (Supplemental Figure 1A, 1D). When we did the same analysis for all the cells where we expressed NES-PABDx2-Spo20 or PASS, we obtained a basal PM/Cyt ratio of 1.1318 ± 0.0954 for NES-PABDx2-Spo20 and a ratio of 0.6861 ± 0.0143 for PASS (Supplemental Figure 1B, 1C, 1E). We believe that the basal localization of these sensors reflects variable PA levels in the PM at resting conditions. FRET based imaging of PA has indicated that there are detectable levels of PA under basal conditions, and this approach also showed some variability in basal PA levels as we see with the spread of Nir1-LNS2’s basal localization (Nishioka et al., 2010). Overall, our data suggests that the high affinity of Nir1-LNS2 for PA is reflected in both its basal localization and its response to stimulations such as PMA.”

    To address the idea that PIP2 is responsible for the basal localization of Nir1-LNS, we have added the following to the discussion on p.15, left column, second paragraph: “Aside from concerns about specificity, the ability of Nir1-LNS2 to interact with PIP2 in liposomes could suggest that the basal PM localization of Nir1-LNS2 is due to it binding PIP2. However, selective depletion of PI(4,5)P2 did not affect basal Nir1-LNS2 localization to the PM (Figure 3C) and was not able to recruit the probe to mitochondria (Figure 4A-C). We did see FKBP-PJ reduce the association of Nir1-LNS2 with the PM under resting conditions (Figure 3E, 3F), suggesting a possible non-specific ionic interaction with polyanionic inositol lipids. Another mechanism to explain these data would be phosphoinositide-dependence of PA production. Phosphoinositides are well-known to regulate the recruitment of PLD isoforms and type II DGKs to the PM as well as their catalytic activity there (Sciorra et al., 2002; Du et al., 2003; Hodgkin et al., 2000; Liscovitch et al., 1994; Kume et al., 2016). Therefore, we suggest that the effects of FKBP-PJ could be reducing basal PLD and DGK activity and hence lowering resting PA levels. That could explain the loss of both basal Nir1-LNS2 PM association when FKBP-PJ is expressed, and Nir1-LNS2’s PM interactions as FKBP-PJ is recruited to the membrane to further deplete phosphoinositides. While this study cannot fully substantiate this hypothesis, the role of PIP2 in PLD activity and PA production is an interesting hypothesis that warrants further investigation.

    #2.3. Fig 1 shows Nir1-LNS2 translocates to plasma membrane upon PMA stimulation in a PLD activity-dependent manner. However, the image in Fig1K is not super convincing since there is already a decent amount of plasma membrane localization of the sensor at t = 0 min, which looks considerably different from the t = 0 min image shown in 1F.

    We have updated the images both in Figure 1F and Figure 1K to best represent the mean basal localization as determined in Supplemental Figure 1.

    #2.4. Fig 3 and Fig 4 need the validation of PIP depletion/production using PIP-binding probes.

    These controls are shown in Figure 4B. We have only included PH-PLCd1 to show PIP2 levels as the large PIP2 production by a PIP5K also indicates the large elevation of the substrate PI4P.

    This control data has now been included in Figure 3, and is referenced by the Figure 3 legend and the following text from p.8, left column, 4th paragraph: “As a negative control, we expressed a doubly catalytically dead mutant of PJ. When PJ-Dead was recruited to the PM, we confirmed that PIP2 and PI4P levels remained unaltered by seeing stable association of the PIP2 biosensor Tubby(c) with the PM (Figure 3A). We observed no loss of the PM localization of Nir1-LNS2 with PJ-Dead recruitment (Figure 3A, 3E). When the active PJ was expressed in HEK293A cells, there was a slight loss of Nir1-LNS2 at the PM even before PJ recruitment (Figure 3B, 3E), although this was not significant as compared to pre-stimulated cells expressing PJ-Dead (Figure 3F). However, Nir1-LNS2 did move off the PM into the cytosol after PJ recruitment, to a similar extent that the PIP2 biosensor Tubby(c) moved off the PM (Figure 3B, 3E). AUC analysis of the Nir1-LNS2 response showed there was a significant reduction of Nir1-LNS2 PM localization (Figure 3G).

            Since PJ depletes both PIP2 and PI4P, we examined which of these lipids specifically contribute to Nir1-LNS2 membrane binding. We utilized an FKBP-INPP5E construct that depletes PIP2 but does not deplete PI4P at the PM, as seen by the significant loss of PM-localized Tubby(c) (Figure 3C). Then FKBP-Sac1, an FKBP-PJ construct that has a catalytically dead INPP5E domain, but an active Sac1 domain was used to deplete PI4P without altering PIP2 levels, as seen by removal of the PI4P biosensor P4Mx1 from the PM (Figure 3D). Recruitment of FKBP-INPP5E did not significantly affect Nir1-LNS2 localization (Figure 3C, 3E, 3G). However, recruitment of FKBP-Sac1 slightly, but not significantly affected Nir1-LNS2 localization (Figure 3D, 3E, 3G). This data suggests that decreasing the anionic charge of the membrane through depletion of PIPs slightly reduces Nir1-LNS2’s ability to interact with the PM, but it doesn’t fully re-localize the sensor. Therefore, this is a small caveat to consider when using Nir1-LNS2 to study PA.”

    #2.“Advance”: The key significance of the manuscript, which is the superiority of Nir1-LNS2 over existing PA-binding probes, is not clear from the data provided. Other than that part, the study does not seem to include significant finding, since the binding of Nir1-LNS2 to PA itself is already known (EMBO Rep. 2013 Oct; 14(10): 891-899, Mol Biol Cell. 2022 Mar 1;33(3):br2).

    While the Kim et al., paper referenced by the reviewer does show that the LNS2 binds to PA, this same group later published data showing that the LNS2 binds to both PA and DAG. (Kim et al., 2015 doi: 10.1016/j.devcel.2015.04.028). Therefore, we believe our data which unequivocally shows that the LNS2 does not bind DAG, is a significant advancement in the field. Aside from the creation of the new biosensor, it progresses our understanding of the mechanism of the Nir family lipid transfer proteins, which are vital to PM lipid homeostasis.

    To highlight this point, we have added the following paragraph to the discussion on p.14, right column, 1st paragraph: “The lack of Nir1-LNS2 binding to DAG-rich liposomes (Figure 2), DAG produced at the mitochondria (Figure 4), and DAG analogs (Figure 5) shows that the LNS2 domains only binds to PA rather than to PA and DAG as has been reported previously (Kim et al., 2015). In this study, we redefined the boundaries of the LNS2 domain based on the structure of the Lipin/Pah family domains and the AlphaFold prediction for the Nir1-LNS2. The new boundaries included the entire fold that is conserved between the Lipins/Pahs and the Nirs. Therefore, we suspect that the expansion of the LNS2 domain in our work explains the differences in our data and the published literature regarding DAG binding. Importantly, the data obtained with our amended Nir1-LNS2 suggests that within the context of the lipid transfer cycle and MCS formation, the Nir family of PITPs translocate to the PM solely based on PA. This information will be important as the field continues to determine the exact mechanism of the Nir PITPs in lipid homeostasis.”

    #3.4. Due to PA's versatile biological roles, the evidence provided by the MCS experiment is far from enough to conclude that Nir1-LNS2 does not interfere with PA function. Further examination of various endogenous pathways is warranted before making the statement "Therefore, Nir1-LNS2 can be used ...... without concern of affecting downstream events".

    We have rewritten the quoted sentence for a more nuanced interpretation on p.13, right column, second paragraph:“While PA has a plethora of cellular functions, the fact that Nir1-LNS2 expression does not disrupt MCS formation shows promise that the high affinity of Nir1-LNS2 will not inhibit downstream PA signaling.”

    #3.5. In Fig. 3A-D (Left), it is unclear to what extent PIPs are reduced after treatment with FKBP-tagged PIP phosphatases. The treatment depicted in the illustration should be accompanied by data, e.g., % of PIPs being degraded after treatment.

    This comment is addressed in our response to #2.4, where we show the addition of control biosensors for PIP2 and PI4P, and also propose new experiments in TIRFM for more sensitive and precise measurements.

    4. Description of analyses that authors prefer not to carry out

    __#1.1a: __Have the authors considered using the DGK inhibitor R59022 to selectively block the conversion of DAG to PA by DGK? Such an experiment could provide additional evidence for the requirement of DGK activity and consequent PA formation for Nir1-LNS2 membrane localization.

    We did indeed attempt experiments with R59022, and have made several unexpected findings with the compound that go way beyond the scope of the current manuscript. In short, although R59022 reduces DGK catalytic activity, it also potently drives over-expressed or endogenous DGKalpha to the plasma membrane, and induces large accumulations of PM PA. This complicated interpretation of data obtained with this compound. We are currently preparing a manuscript detailing the novel and unexpected effects.

    #3.6. In Fig. 4C, the plasma membrane (PM) localization of Nir1-LNS2 and NES-PABDx2-Spo20, as determined by the "intensity PM/Cyto," should be analyzed following the ectopic production of PI4P and PIP2. Although mitochondria do not apparently recruit Nir1-LNS2 or NES-PABDx2-Spo20 after induced PI4P and PIP2 production, it remains possible that the subsequent trafficking of PI4P and PIP2 from mitochondria might sequester the biosensors away from the PM into the cytoplasm, thereby reducing the "intensity PM/Cyto" of Nir1-LNS2.

    We cannot easily determine the PM/cyt ratio in this experiment as we included a mitochondrial marker rather than a PM marker when imaging. However, based on the images, there is no change in the PM intensity of the Nir1-LNS2 and NES-PABDx2-Spo20 biosensors. The images included in Figure 4 are representative of this localization.

    #3.7. It would be valuable to determine the half-life (stability) of Nir1-LNS2.

    In all of our transient transfections, the Nir1-LNS2 shows good stability where we don’t expect degradation to be a major concern. Furthermore, stability has not usually been factor considered in the creation any of the current widely used lipid biosensors.

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    Referee #3

    Evidence, reproducibility and clarity

    Weckerly et al. introduced a fluorescently tagged Nir1-LNS2 construct capable of binding to both PA and PIP2 in vitro, yet selectively targeting PA-enriched membranes in cells. Their findings demonstrate that Nir1-LNS2 exhibits heightened responsiveness to PA, allowing the authors to uncover a modest contribution of PLD to PA production downstream of muscarinic receptors, a phenomenon not visualized with previous Spo20-based biosensors. Thus, Nir1-LNS2 is a sensitive biosensor, potentially providing researchers with a powerful new tool for real-time investigation of PA dynamics in live cells. The manuscript is well-written, with major conclusions supported by experimental evidence. The tool developed in this study holds significant importance for the field of lipid biology. However, missing controls and weaknesses from the in vitro analysis reduce the overall impact of this work. The authors are encouraged to address the following comments to further strengthen their conclusions:

    Major Points:

    1. The direct measurement of the binding affinity of Nir1-LNS2 with PA, e.g., Kd, is essential; this information will help the field explore the potential usage of Nir1-LNS2.
    2. As mentioned by the authors, there is a confusing inconsistency regarding why Nir1-LNS2 binds to PIP2 in vitro but not in cells. Going beyond what has been discussed in the manuscript, there is a possibility that PIP2 could induce Nir1-LNS2 aggregation, leading to pelleting after centrifugation, among many other possibilities. I recommend the authors perform additional in vitro experiments, including but not limited to the liposome floatation assay to directly examine Nir1-LNS2 binding to the liposomes with varied compositions.
    3. In Fig. 2D, it would be beneficial to examine the constructs Nir1-613-630 and Nir1-631-894, comparing them with Nir1-LNS2 using liposome sedimentation and floatation assays to evaluate the contribution of the SIDGS motif and the amphipathic helix in binding PA.
    4. Due to PA's versatile biological roles, the evidence provided by the MCS experiment is far from enough to conclude that Nir1-LNS2 does not interfere with PA function. Further examination of various endogenous pathways is warranted before making the statement "Therefore, Nir1-LNS2 can be used ...... without concern of affecting downstream events".

    Minor Points:

    1. In Fig. 3A-D (Left), it is unclear to what extent PIPs are reduced after treatment with FKBP-tagged PIP phosphatases. The treatment depicted in the illustration should be accompanied by data, e.g., % of PIPs being degraded after treatment.
    2. In Fig. 4C, the plasma membrane (PM) localization of Nir1-LNS2 and NES-PABDx2-Spo20, as determined by the "intensity PM/Cyto," should be analyzed following the ectopic production of PI4P and PIP2. Although mitochondria do not apparently recruit Nir1-LNS2 or NES-PABDx2-Spo20 after induced PI4P and PIP2 production, it remains possible that the subsequent trafficking of PI4P and PIP2 from mitochondria might sequester the biosensors away from the PM into the cytoplasm, thereby reducing the "intensity PM/Cyto" of Nir1-LNS2.
    3. It would be valuable to determine the half-life (stability) of Nir1-LNS2.

    Significance

    Tracking intracellular phosphatidic acid (PA) in live cells is essential for understanding its cellular functions, leading to the development of genetically encoded lipid biosensors. While several PA biosensors have been developed, they often suffer from limited sensitivity or specificity.

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    Referee #2

    Evidence, reproducibility and clarity

    Summary:

    The authors reported a new PA-binding probe Nir1-LNS2, which potentially offers advantages over conventional tools with its higher sensitivity for PA. The authors performed extensive characterization in different cell lines to test the ability of Nir1-LNS2 to selectively bind to PA without disrupting endogenous PA signaling. While the tool is potentially useful as a new PA-binding probe with higher spatiotemporal precision, the data provided in the manuscript are not enough to support their claims and conclusions. Especially, the data do not fully support that the Nir1-LNS2 offers more sensitive and selective binding to PA than conventional PA-binding probes using Spo20.

    Major comments:

    1. Nir1-LNS2 seems to show variable basal localization across different representative images presented in the manuscript. A part of them were justified by the effect of other anionic species by PIP (such as Fig 3 where they co-expressed PIP-degrading enzymes). For example, cells in Fig 1F and those in Fig 4F show quite different basal localization of Nir1-LNS2. Is it due to difference in expression level, cell conditions, or other factors? The significant amount of plasma membrane basal localization seems to indicate that Nir1-LNS2 localization is affected by its binding to PI(4,5)P2. The significant and potentially variable plasma membrane localization of Nir1-LNS1 can limit the utility of this probe.
    2. The authors mention high affinity of Nir1-LNS2, but it lacks in vitro characterization that should demonstrate the higher affinity of Nir1-LNS2 compared to conventional probes such as Spo20. The authors should perform side-by-side comparison in Fig 2 to compare the PA affinity and specificity of Nir1-LNS2 compared to Spo20.

    Minor comments:

    1. Fig 1 shows Nir1-LNS2 translocates to plasma membrane upon PMA stimulation in a PLD activity-dependent manner. However, the image in Fig1K is not super convincing since there is already a decent amount of plasma membrane localization of the sensor at t = 0 min, which looks considereably different from the t = 0 min image shown in 1F.
    2. Fig 3 and Fig 4 need the validation of PIP depletion/production using PIP-binding probes.
    3. In Discussion: "while in vivo it solely binds to PA (Fig 4)" - this claim does not seem to be true according to Fig 4, where the overexpression of PIP-degrading enzymes did affect the Nir1-LNS2 basal localization.

    Significance

    General assessment:

    The existing PA-binding probe using Spo20 is indeed quite blunt, which takes minutes to see appreciable accumulation of the probe upon PA production. Nir1-LNS2 can be indeed useful if it offers better spatiotemporal precision. However, the advantage of this tool over existing tools is not convincing without head-to-head comparison of either (1) in vitro characterization of PA binding affinity and selectivity between Nir1-LNS2 and Spo20 or (2) response to PA produced on different subcellular localizations other than plasma membrane and mitochondria (e.g., endosomes, golgi, and endoplasmic reticulum).

    Advance:

    The key significance of the manuscript, which is the superiority of Nir1-LNS2 over existing PA-binding probes, is not clear from the data provided. Other than that part, the study does not seem to include significant finding, since the binding of Nir1-LNS2 to PA itself is already known (EMBO Rep. 2013 Oct; 14(10): 891-899, Mol Biol Cell. 2022 Mar 1;33(3):br2).

    Audience:

    The lipid biology community would be highly interested in using the new PA-binding tool to study lipid localization in live cells.

    My expertise is PA signaling and deveopment of engineered phospholipase Ds, which can produce PA on demand at various subcellular locations.

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    Referee #1

    Evidence, reproducibility and clarity

    Summary:

    The authors designed, created, and validated a fluorescently tagged sensor protein that binds with high affinity to the signaling phospholipid PA in cells. The LNS2 PA-binding domain used originates from the lipid transfer protein Nir1, and shares conserved features with lipins. The novel sensor outperforms the commonly used Spo20-based PA probes, although it also suffers from binding to PIP2 in vitro (liposomes) and from PIPs affecting its membrane binding in vivo. Importantly, the authors demonstrate that PA but not DAG or PIP2 is sufficient for membrane binding of Nir1-LNS2 in cells, validating Nir1-LNS2 as a PA-sensor in fluorescence microscopy studies.

    Major comments:

    1. CCH treatment of HEK293A cells leads to the PM localization of the DAG sensor C1ab-Prkd1 as well as Nir1-LNS2 (Fig. 5), and the kinetics of these changes - Nir1-LNS2 would lag behind C1ab-Prkd1 fluorescence - is taken as evidence for Nir1-LNS2's specific binding of PA rather than DAG: Pag. 10: 'When we look at the first 2-minutes after CCh addition, we see that C1ab-Prkd1 moves to the PM much faster than Nir1-LNS2 does (Figure 5D). The delay in Nir1-LNS2 translocation makes sense given DAG is produced first and then converted into PA, again indicating that Nir1-LNS2 is specific for PA.' Fig. 5 legend: 'The Nir1-LNS2 response to PLC depends on PA and not DAG.(...) (D) Nir1-LNS2 translocation to the PM (data replicated from Figure 5C) lags behind C1ab-Prkd1 (data replicated from Figure 5B) in response to CCh addition.' The validity of the conclusion from this experiment seems questionable. The argument relies on the low values of Nir1-LNS2's normalized fluorescence intensity compared to C1ab-Prkd1's, in the first two minutes of stimulation, when DAG is expected to accumulate (Fig. 5D). However, if Nir1-LNS2 would bind DAG, the resulting fluorescence is expected to be low, i.e. compared to the much higher signal resulting from subsequent PA binding. Moreover, in this system, which co-expresses the two sensor proteins, competition in binding may account for the apparent precedence of one sensor over the other. Thus, even if the increase in C1ab-Prkd1 fluorescence would precede Nir1-LNS2 - following the authors' interpretation - this would not exclude binding of DAG by Nir1-LNS2. Fig. 5D shows the confocal images of cells after 30 sec CCH treatment using the two sensors, next to the respective controls, replicated from Fig. 5B/C. However, in Fig. 5D, different colors are used for Nir1-LNS2 than in Fig. 5C, which makes comparison along the time course difficult. In conclusion, the data presented in Fig. 5 do not exclude DAG binding by Nir1-LNS2, and modification of the conclusions from this experiment throughout the ms (including the cited sentences) is recommended. Consider removal of Figure 5D. Despite these considerations, the authors' final conclusion regarding the specificity of Nir1-LNS2 towards PA appears well supported (e.g. by the data presented in Figure 4).

    Have the authors considered using the DGK inhibitor R59022 to selectively block the conversion of DAG to PA by DGK? Such an experiment could provide additional evidence for the requirement of DGK activity and consequent PA formation for Nir1-LNS2 membrane localization.

    1. P 4 and Fig. 2 mention a 'novel domain structure', responsible for binding of PA and PIP2 (in vitro). What exactly is this novel domain structure? Why have the two parts of Nir1-LNS, the AAH and the 263 amino acid domain, not been tested in similar liposome assays as in Fig. 2A? Lipid binding in vivo, as tested in the experiment of Fig. 2D, is confounded by endogenous PA binding proteins. PA is expected to bind the SIDGS motif, as this is conserved from the Lipin catalytic motif (p. 5). However, experimental evidence of this appears to be lacking. Nevertheless, it is several times presented as a fact in the text. Pag. 7: 'This suggests that the SIDGS motif alone is not the sole PA binding pocket as the LNS2 domain requires both that motif and the amphipathic helix for sustained binding to membrane embedded PA.' Pag. 13: '... One reason for this difference could be that the Nir1-LNS2 is not a novel bona fide PA binding pocket. Rather, it requires both the SIDGS motif and an N-terminal amphipathic helix for membrane interactions (Figure 2). These conclusions therefore appear to be not accurate and would need te be rephrased.

    If the authors could show that PA binding by Nir1-LNS can be eliminated by mutating residues in the SIDGS motif, this would not only substantiate the above claims, but also make for a negative control protein, next to Nir1-LNS2. For future applications of Nir1-LNS2 as PA biosensor in other organisms this would be useful.

    1. Pag. 7: '...small caveat to consider when using Nir1-LNS2 to study PA, the data also demonstrates that Nir1-LNS2 is not specifically interacting with any of the PM PIPs in cellular membranes.' This seems not accurate, since the data in Fig. 3 suggest that PI4P could be involved in membrane localization of Nir1-LNS2. It remains however unresolved whether this is a specific interaction with this PIP.
    2. Please note that the presence of PE increases the ionization (and negative charge) of PIP2 (Graber et al., 2012) rather than dilutes the negative charge as stated in the Discussion on p.13. Please revise!

    Minor comments:

    1. P 1 The authors may consider adding a 4th criterium for a lipid biosensor: the sensor should not serve as a sink for the lipid by removing/sequestering it from the active pool, thereby interfering with other interactions/conversions.
    2. Nir1 lacks a PITP domain (Fig. 1), yet is referred to as lipid transfer protein: please elaborate/explain.
    3. Fig. 2A suggests cooperativity in binding of Nir1-LNS2 to PA-containing liposomes. Please mention/comment! Does binding to PIP2-containing liposomes also exhibit cooperativity?
    4. Indicate the concentrations of PC and protein in the legend to Fig. 2 panels A and B. M&M says 2 mM PC, according to the PA-concentrations above panel 2A, this should be 1 mM. Please clarify.
    5. In Fig. 2B, PI is missing. Any specific reason?
    6. Move Fig. 2C to the Introduction and extend it to illustrate the shared conserved features of Nir1-LNS2 and lipin.
    7. P 13. 'While real-time IMPACT does not directly report on PA levels as it does not use the endogenous PLD substrate PC, ...' It is true that this method doesn't directly report on PA levels, but that is because it uses a click chemistry probe as substrate for PLD's transphosphatidylation reaction. Contrary though to what is stated by the authors, this reaction still uses the endogenous substrate of PLD, PC (Liang et al. 2019; www.pnas.org/cgi/doi/10.1073/pnas.1903949116).
    8. Fig. 1K: Control must have been treated with PMA plus vehicle (DMSO); if so, please indicate that vehicle was added.
    9. Figure 6C: How is Ft/Fpre defined? Add to legend.
    10. P 4: 'The boundaries of the Nir2-LNS2 (Uniprot: O00562) and Nir3-LNS2 (Uniprot: Q9BZ72) were also defined using AlphaFold predictions of the structure of these domains.' How was the extent of each of these domains determined - they are much larger than the previously published sequences of LNS2 domains (Kim et al. 2013; Embo Rep. 14:891-899. doi:10.1038/embor.2013.113)?
    11. In Fig. 3 legend, specify the starting condition of Nir1-LNS2 binding?? Which fluorescence are we looking at?
    12. In legend to Fig. 6 please specify the fluorescent tags used. Have they been shown not to affect protein function?

    Significance

    The development of a new, superior PA-sensor is a significant advance in the fields of lipid signaling and specific lipid-protein interactions, that will benefit research on lipid-mediated cellular signaling and intracellular lipid trafficking.

    This reviewer's expertise encompasses lipid metabolism and lipid-protein interactions, not so much fluorescence microscopy.

  5. Version published to 10.1101/2024.02.28.582557v1 on bioRxiv