Improved ANAP incorporation and VCF analysis reveal details of P2X7 current facilitation and a limited conformational interplay between ATP binding and the intracellular ballast domain

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Version of Record published: January 20, 2023 (This version)
Accepted Manuscript published: January 4, 2023 (Go to version)
Accepted: January 3, 2023
Preprint posted: September 1, 2022 (Go to version)
Received: August 5, 2022

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Drug Discovery: Decoding the mechanisms of allostery

Saif Khan, Cornelius Gati

Insight Jun 2, 2023
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Abstract

The large intracellular C-terminus of the pro-inflammatory P2X7 ion channel receptor (P2X7R) is associated with diverse P2X7R-specific functions. Cryo-EM structures of the closed and ATP-bound open full-length P2X7R recently identified a membrane-associated anchoring domain, an open-state stabilizing “cap” domain, and a globular “ballast domain” containing GTP/GDP and dinuclear Zn²⁺-binding sites with unknown functions. To investigate protein dynamics during channel activation, we improved incorporation of the environment-sensitive fluorescent unnatural amino acid L-3-(6-acetylnaphthalen-2-ylamino)–2-aminopropanoic acid (ANAP) into Xenopus laevis oocyte-expressed P2X7Rs and performed voltage clamp fluorometry. While we confirmed predicted conformational changes within the extracellular and the transmembrane domains, only 3 out of 41 mutants containing ANAP in the C-terminal domain resulted in ATP-induced fluorescence changes. We conclude that the ballast domain functions rather independently from the extracellular ATP binding domain and might require activation by additional ligands and/or protein interactions. Novel tools to study these are presented.

Editor's evaluation

This manuscript constitutes a valuable foray into the conformational rearrangements throughout various domains of the notoriously difficult-to-study P2X7 receptor, with a focus on the enigmatic intracellular 'ballast' domain. The molecular origin of the facilitation process and effects by intracellular factors will require future study, but the authors provide convincing evidence that the ballast domain is unlikely to undergo major conformational changes upon ATP-induced gating. The work is of interest to those interested in the role of enzymatically active intracellular domains of membrane proteins.

https://doi.org/10.7554/eLife.82479.sa0

Introduction

P2X receptors (P2XR) are trimeric non-selective cation channels that are activated by extracellular adenosine triphosphate (ATP). The structure of a single P2X subunit has been compared to a dolphin, with two transmembrane domains (TM1 and TM2) that form the ‘fluke’, and a large extracellular domain, comprising the ‘body’, left and right ‘flippers’, and a ‘head’ domain that reaches over one of three inter-subunit ATP binding sites (Kawate et al., 2009). The intracellular N- and C-termini are short in most P2X subtypes and have only been resolved in the open state of the P2X3R and, more recently, in the open and closed states of the P2X7R (Mansoor et al., 2016; McCarthy et al., 2019). The pro-inflammatory P2X7 subtype is expressed in immune cells and considered an important drug target. In contrast to the other P2XR family members, it has a low ATP sensitivity, shows complete lack of desensitization, and contains a large intracellular C-terminus (240 amino acids [aa]), which mediates diverse downstream effects such as interleukin secretion, plasma membrane permeabilization, blebbing, phosphatidylserine flip, and cell death (Kopp et al., 2019). The recently determined cryo-EM structures of the full-length rat P2X7R in the apo/closed and ATP-bound open states (McCarthy et al., 2019) did not only elucidate details of P2X desensitization, but finally unveiled the structure of the large P2X7 C-terminus. Accordingly, intertwined β-strands from all three subunits form an open state-stabilizing ‘cap domain’, that was also found in the P2X3R (Mansoor et al., 2016). In the P2X7R, however, this ‘cap’ is stabilized by a highly palmitoylated membrane-associated ‘Cys-anchor’ domain, which prevents desensitization. The remaining residues 393–595 fold into a dense globular structure (the so-called ‘ballast domain’), which contains a novel guanosine nucleotide binding motif and a dinuclear zinc binding site. A stretch of 27–29 aa (S443-R471) was not resolved, and it is unclear if each ballast domain is formed by a single subunit or if a domain swap occurs between subunits (McCarthy et al., 2019). While these structures represent a milestone in P2X7 research, the transition dynamics between receptor states in a cellular environment as well as the molecular function of the ballast domain and how it is affected by ATP binding remain unclear. Likewise, the molecular mechanism of current facilitation, a P2X7-characteristic process that describes faster and/or increased current responses upon repeated ATP application, is not understood. In this study, we set out to determine conformational changes associated with P2X7-specific functions by voltage clamp fluorometry (VCF). This method allows simultaneous recording of current responses and associated molecular movements that are reported by an environment-sensitive fluorophore. We have previously used site-specific cysteine-substitution and the thiol-reactive fluorophore tetramethyl-rhodamine-maleimide (TMRM) to show a closing movement of the head domain during activation of the oocyte-expressed P2X1R (Lörinczi et al., 2012). However, this procedure is limited to extracellularly accessible residues. To investigate intracellular rearrangements, we therefore employed the fluorescent unnatural amino acid (fUAA) L-3-(6-acetylnaphthalen-2-ylamino)–2-aminopropanoic acid (ANAP) (Lee et al., 2009). This can be site-specifically incorporated into a protein by repurposing the amber stop codon (TAG) and introducing a corresponding suppressor tRNA (CUA anticodon) loaded with ANAP. A plasmid encoding an ANAP-specific bio-orthogonal suppressor tRNA/aminoacyl-tRNA synthetase pair (Chatterjee et al., 2013) has been obtained by co-evolution and selection (Lee et al., 2009) and was successfully used to study voltage-gated and ligand-gated ion channels (Andriani and Kubo, 2021; Kalstrup and Blunck, 2018; Kalstrup and Blunck, 2013; Soh et al., 2017; Wulf and Pless, 2018). This stop-codon suppression can, however, lead to premature translational termination or aberrant stop-codon substitution (read-through) (Braun et al., 2020; Kalstrup and Blunck, 2017; Klippenstein et al., 2018; Pless et al., 2015; Poulsen et al., 2019).

Here, we provide an improved method for fUAA incorporation into Xenopus laevis oocyte-expressed proteins and analyzed membrane surface expression and functionality for a total of 61 P2X7R mutants with ANAP substitutions in the extracellular head domain, the second transmembrane domain (TM2), and the intracellular N- and C-termini. Using VCF, we identified 19 positions in which ANAP reported ATP-induced localized rearrangements. To further expand the VCF toolbox, we demonstrate simultaneous recordings of fluorescence changes from ANAP in combination with other fluorophores. We conclude from our data that (i) current facilitation is intrinsic to the P2X7 protein and likely caused by a change in gating and (ii) the cytoplasmic ballast functions rather independently from the extracellular ligand binding domain and might require activation by additional ligands or protein interactions.

Results

Improved ANAP incorporation by cytosolic co-injection of mutated X. laevis eRF1 cRNA

To implement and optimize a protocol for incorporation of ANAP into Xenopus oocyte-expressed protein, we initially used the P2X1R as it was already intensively studied in our lab (Lörinczi et al., 2012) and has functional similarity with the P2X3R, which at the beginning of this study, represented the only P2XR for which the intracellular termini were resolved (Mansoor et al., 2016). Using the original 2-step-injection protocol (Kalstrup and Blunck, 2017; Kalstrup and Blunck, 2013) and a simplified procedure where all components required for the expression of UAA-containing receptors are injected simultaneously (Figure 1A and C), we introduced ANAP into non-conserved positions within the N-terminally His-tagged P2X1R N- and C-termini (position 10 and 388, respectively, ANAP substitutions indicated by *) and compared the formation of full-length and truncated receptors in the plasma membrane by SDS-PAGE. As seen in Figure 1B, ANAP-containing P2X1Rs were efficiently expressed and virtually no read-through product was detected in the absence of ANAP. The new protocol resulted in less variable protein expression but also a reduced ratio of full-length and truncated His-P2X1 EGFP protein (Figure 1D). The relative amount of full-length protein was neither increased by different forms of ANAP application nor by variation of injection protocols (Figure 1—figure supplement 1A and B). Therefore, we tested if a mutated eukaryotic release factor (eRF1(E55D)), which was previously shown to favor UAA-incorporation over translational termination in HEK293T cells (Gordon et al., 2018; Schmied et al., 2014) could also be used in the Xenopus oocyte expression system. Indeed, co-injection of either purified X. laevis eRF1(E55D) protein (Figure 1—figure supplement 1B) or the respective in vitro synthesized cRNA (Figure 1C and D) resulted in a more than threefold higher ratio of full-length and truncated receptor constructs compared to the 1-step injection method without eRF1(E55D) and a smaller standard deviation compared to the 2-step injection method (1-step+eRF1(E55D): 1.469±0.229; 1-step: 0.418±0.082; 2-step: 1.603±0.933; mean ± S.D.). The applicability of this approach was confirmed for the hα1 glycine receptor (GlyR) A52* mutant (Figure 1—figure supplement 1C; Soh et al., 2017). In conclusion, this optimized protocol led to more reproducible expression and increased formation of full-length ANAP-labeled receptors and was used in all following experiments.

Figure 1 with 1 supplement see all

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Optimization of fluorescent unnatural amino acid (fUAA) incorporation into *Xenopus laevis* oocyte-expressed P2X1 receptor.

(A) Molecular structure of L-3-(6-acetylnaphthalen-2-ylamino)–2-aminopropanoic acid (ANAP) and schematic representation of the 2-step injection method for site-specific ANAP incorporation using the *amber* stop-codon (UAG) and a plasmid containing the orthogonal tRNA/tRNA-synthetase pair. (B) Representative SDS-PAGE analysis of plasma membrane-expressed ANAP-labeled (S10* or S388*) rat P2X1Rs (46 kDa without glycosylation). A C-terminal EGFP-tag (27 kDa) was added as indicated to enable detection of premature termination at position 388. Oocytes were injected as shown in A and labeled with membrane impermeable Cy5-NHS ester. His-tagged P2X1Rs were extracted in 0.5% n-dodecyl-β-D-maltoside, purified via Ni²⁺-NTA agarose, and separated by SDS-PAGE (8%). Noninjected oocytes and oocytes injected only with the plasmid pANAP, P2X1 cRNA without the *amber* stop codon (Wt), or without ANAP (as indicated) served as controls. Note, that twice the amount of protein was loaded for P2X1(S10*). Ø indicates empty lanes. Two to three independent experiments were performed. (C) Representation of the 1-step injection method and all components required for UAA-labeling plus optional *X. laevis* eRF1(E55D) cRNA (left) and (right) scheme of protein translation termination by eRF1 (upper panel) and how overexpression of the mutated form of eRF1 favors *amber*-encoded fUAA incorporation by outcompeting endogenous eRF1 (lower panel). (D) Comparison of Cy5-labeled membrane-expressed full-length and truncated His-rP2X1-EGFP(388*) ratios upon expression by the 2-step and 1-step injection method with or without eRF1(E55D) co-expression. A representative SDS-PAGE gel (prepared as in B) and statistical analysis of data from 6 to 11 experiments including oocytes from 4 to 6 different *X. laevis* frogs per group are shown. Data are represented as mean ± S.D., and significance was determined by a two-tailed unpaired Welch’s t-test and is indicated as *p<0.05 and ****p<0.0001.

Figure 1—source data 1 Original gel, Figure 1B.: https://cdn.elifesciences.org/articles/82479/elife-82479-fig1-data1-v2.zip
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Figure 1—source data 2 Original gel, Figure 1D.: https://cdn.elifesciences.org/articles/82479/elife-82479-fig1-data2-v2.zip
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Figure 1—source data 3 Original gels for bar graph in Figure 1D: Gel2, Gel3, Gel4, and Gel5.: https://cdn.elifesciences.org/articles/82479/elife-82479-fig1-data3-v2.zip
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Figure 1—source data 4 Original gels for bar graph in Figure 1D: GelA, GelB, and GelC.: https://cdn.elifesciences.org/articles/82479/elife-82479-fig1-data4-v2.zip
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Evaluation of plasma membrane expression of full-length ANAP-containing P2X7Rs

Next, we incorporated ANAP into the P2X7R in sites chosen based on previous structure-function studies and the cryo-EM structures (McCarthy et al., 2019). As a positive control, we first introduced ANAP into the head domain (Figure 2A and B), which is known to undergo substantial movements and/or ligand interactions with clear changes of TMRM fluorescence in the P2X1R (Lörinczi et al., 2012) and P2X7R (Figure 2—figure supplement 1A). Next, based on the comparison of the P2X4 and P2X3 crystal structures in the open and closed states (Hattori and Gouaux, 2012; Kawate et al., 2009; Mansoor et al., 2016), and the identification of the human P2X7 channel gate and selectivity filter around residue S342 (Pippel et al., 2017), we selected positions in the second transmembrane helix. Finally, we introduced ANAP throughout the intracellular region in positions that we suspected to undergo conformational changes upon channel activation, as well as in six positions in the unresolved 29 aa stretch. As shown in Figure 2C, all constructs with ANAP substitutions in the N-terminus and the head domain as well as three out of four constructs with substitutions in TM2 were formed in full length, indicating that receptors that are truncated before or within TM2 are retained in the endoplasmic reticulum and likely undergo degradation. Interestingly, ANAP incorporation into G338 completely prevented membrane incorporation while cysteine substitution in the equivalent position of human P2X7R led previously to surface-expressed, but non-functional receptors (Pippel et al., 2017).

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P2X7Rs containing L-3-(6-acetylnaphthalen-2-ylamino)–2-aminopropanoic acid (ANAP) at various positions within the extracellular, transmembrane, and cytoplasmic regions are expressed in the plasma membrane and functional.

(**A, B**) Surface representations of the rat P2X7 Cryo-EM structure in the open state (PDB ID: 6u9w). The different domains (A) and selected sites of ANAP substitutions (B) are indicated in one subunit while the two other subunits are shown in gray and wheat, respectively. (C) Evaluation of surface expression and functionality of P2X7Rs generated from constructs containing an *amber* stop codon in the indicated positions. *X. laevis* oocytes expressing the constructs were labeled with membrane-impermeant Cy5-NHS ester. His-tagged P2X7Rs were extracted in 0.5% n-dodecyl-β-D-maltoside, purified via Ni²⁺ NTA agarose, and analyzed by SDS-PAGE (8%). Symbols indicate current responses to 0.3 mM ATP as determined by two-electrode voltage clamp recordings in the voltage clamp fluorometry setup: +, functional and currents comparable to wt P2X7 after 2–4 days of expression; (+), functional and currents comparable to wt P2X7 after 5–7 days of expression; –, not functional or currents ≤0.5 μA and not reproducible after 4 days. Representative data from two to five independent biochemical experiments are shown.

Figure 2—source data 1 Original gel (1), Figure 2C (N-terminus).: https://cdn.elifesciences.org/articles/82479/elife-82479-fig2-data1-v2.zip
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Figure 2—source data 2 Original gel (2), Figure 2C (head domain and TM2).: https://cdn.elifesciences.org/articles/82479/elife-82479-fig2-data2-v2.zip
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Figure 2—source data 3 Original gel (3), Figure 2C (juxtamembrane).: https://cdn.elifesciences.org/articles/82479/elife-82479-fig2-data3-v2.zip
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Figure 2—source data 4 Original gel (4), Figure 2C (anchor, cap, and ballast domain).: https://cdn.elifesciences.org/articles/82479/elife-82479-fig2-data4-v2.zip
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Figure 2—source data 5 Original gel (5), Figure 2C (unresolved).: https://cdn.elifesciences.org/articles/82479/elife-82479-fig2-data5-v2.zip
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Figure 2—source data 6 Original gel (6), Figure 2C (ballast domain, part a).: https://cdn.elifesciences.org/articles/82479/elife-82479-fig2-data6-v2.zip
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Figure 2—source data 7 Original gel (7), Figure 2C (ballast domain, part b).: https://cdn.elifesciences.org/articles/82479/elife-82479-fig2-data7-v2.zip
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Starting from T357 in the C-terminus, introduction of the amber stop codon resulted in variable ratios of truncated and full-length receptors. Surface expression of full-length receptors was particularly low for constructs containing ANAP in the C-terminal cap (K387*, C388*) and ballast (I577*) domains, while it was most efficient for ANAP-substitutions in positions 517–537 (in particular L527* and E537*) and in the very C-terminus (Y595* and 596*).

In summary, most substitutions within the C-terminus led to a dominant formation of truncated P2X7 protein besides full-length receptors. Nevertheless, the majority of these constructs showed clear current responses (Figure 2C, Table 1). Since the truncated forms were not expected to interfere with the fluorescence signal, functional constructs that were expressed at least partly in full length were further analyzed by VCF.

Table 1

Summary of surface expression, current responses (ΔI), and L-3-(6-acetylnaphthalen-2-ylamino)–2-aminopropanoic acid fluorescence changes (%ΔF/F) of the investigated P2X7 mutants.

	Position	Surface expression		ΔI	%ΔF/F Filter set 1		%ΔF/F Filter set 2
	Position	Full-length	Truncated	ΔI	430–470 nm	470–500 nm	430–490 nm	>500 nm
N-terminus	A3	+	–	+	↑	↑	↑	↑
	C5	+	–	+	(–)	(–)	↑	↑
	S6	+	–	–	(–)	(–)	(–)	(–)
	W7	+	–	+	(↑)	(↑)	↑	↑
	V10	+	–	(+)	(–)	(–)	(↑)	(↑)
	F11	+	–	+	(–)	(–)	↑	↑
	K17	+	–	(+)	(–)	(–)	(–)	(–)
Head domain	P120	+	–	+	(↑)	(↑)	↑	↑
	E121	+	–	+	(–)	(–)	↑	↑
	Y122	+	–	+	↓	↓	↓	↓
	P123	+	–	+	n.d.	n.d.	↓	↓
	S124	+	–	+	↑	↑	↑	↓
	R125	+	–	+	↑	↑	↑	↓
	G126	+	–	+	↑	↑	↑	↑
	K127	+	–	+	↓	↓	↓	↓
	Q128	+	–	+	–	–	↑	↓
TM2	G338	–	–	–	n.d.	n.d.	(–)	(–)
	S339	+	–	+	↑	↑	↑	↑
	T340	+	–	+	– / ↓	↑	↑	↑
	L341	+	–	+	(↑)	(↑)	↑	↑
C-terminus	N356	+	–	(+)	(–)	(–)	(–)	(–)
	T357	+	+	(+)	(–)	(–)	(–)	(–)
	Y358	+	+	+	(–)	(–)	–	–
	A359	+	+	(+)	(–)	(–)	(–)	(–)
	T361	+	+	+	(–)	(↓)	–	↓
	R364	+	+	(+)	n.d.	n.d.	(–)	(–)
	C371	+	+	+	–	–	(–)	(–)
	A378	+	+	(+)	(–)	(–)	n.d.	n.d.
	R385	+	+	(+)	n.d.	n.d.	(–)	(–)
	K387	+	+	–	n.d.	n.d.	(–)	(–)
	C388	+	+	–	n.d.	n.d.	(–)	(–)
	V392	+	+	(+)	(–)	(–)	(–)	(–)
	E406	+	+	(+)	(–)	(–)	(–)	(–)
	Q422	+	+	–	n.d.	n.d.	(–)	(–)
	D423	+	+	+	(–)	(–)	↑	↑
	V424	+	+	–	n.d.	n.d.	(–)	(–)
	Q433	+	+	(+)	(–)	(–)	n.d.	n.d.
	T434	+	+	(+)	(–)	(–)	n.d.	n.d.
	F436	+	+	+	n.d.	n.d.	(–)	(–)
Unresolved	S445	+	+	+	(–)	(–)	–	(–)
	P450	+	+	+	–	–	–	–
	Q455	+	+	+	–	–	–	–
	Q460	+	+	(+)	(–)	(–)	(–)	(–)
	E465	+	+	(+)	(–)	(–)	n.d.	n.d.
	S470	+	+	(+)	(–)	(–)	n.d.	n.d.
C-terminus	E489	+	+	+	(–)	(–)	–	–
	N490	+	+	+	(–)	(–)	–	–
	V517	+	+	+	n.d.	n.d.	–	–
	L523	+	+	+	n.d.	n.d.	–	–
	L527	+	+	+	n.d.	n.d.	–	–
	L536	+	+	+	(–)	(–)	–	–
	E537	+	+	+	–	–	–	–
	G538	–	+	–	(–)	(–)	(–)	(–)
	E539	+	+	+	(–)	(–)	–	–
	A564	+	?	+	n.d.	n.d.	↑	↑
	L569	+	?	(+)	n.d.	n.d.	(–)	(–)
	I577	+	?	–	n.d.	n.d.	(–)	(–)
	Q585	+	?	+	(–)	(–)	–	–
	G586	+	?	+	(–)	(–)	–	–
	Y595	+	?	+	–	–	–	–
	596	+	?	+	–	–	–	–

+ and − indicate presence and absence of protein or signals, respectively. In case of current responses, + means response comparable to wt receptors and (+) means reduced responses. ↑ and ↓ indicate positive and negative fluorescence signals, respectively. 3–50 oocytes were measured per construct and filter set. In case of fluorescence responses, symbols in brackets indicate where less than three recordings met the criteria defined in the methods (mostly because of impaired fucntionality) and represent tendencies only. ?, not distinguishable (because of similar length of full-length and truncated constructs); n.d., not determined.

Table 1—source data 1 Summarized data for Table 1 with assignment to the original VCF recordings; also including data from Figure 2—figure supplement 1B (box plot); Figure 3C, D, E, F; Figure 3—figure supplement 3C; Figure 4B, C, D; Figure 4—figure supplement 1B; and Figure 5B, C, D. The respective original recordings are deposited with Dryad.: https://cdn.elifesciences.org/articles/82479/elife-82479-table1-data1-v2.zip
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Recording of ANAP fluorescence in the head domain reveals mainly gating-associated movements

Next, we recorded ANAP fluorescence changes upon application of 0.3 mM ATP. Control oocytes expressing wt P2X7R showed a gradual fluorescence decrease during ATP application, even when no ANAP was injected (Figure 3—figure supplement 1). A similar fluorescence drift was observed with the P2X2R, but not with the hɑ1 GlyR or the fast-desensitizing P2X1R. The reason for this drift is unclear but needs to be considered when evaluating mutants with small negative fluorescence changes. Specificity of tRNA loading and ANAP-incorporation was evaluated in further control experiments (Figure 3—figure supplement 2). Only recordings that met specific inclusion criteria (see Methods) were considered for analysis.

As a prodan derivative, ANAP is highly sensitive to the polarity of its environment and shows a redshift in emission with increasing polarity (Lee et al., 2009; Weber and Farris, 1979). Consequently, alterations in ANAP fluorescence can be attributed to (i) quenching by ligands, small molecules, or (aromatic) aa, (ii) spectral shift due to changes in the polarity of the environment, or (iii) a combination of these two effects, e.g., in case of ligand interaction.

To allow differentiation between presumably wavelength-independent (de-)quenching of ANAP fluorescence by other molecules or spectral shifts due to changes in the polarity, we simultaneously recorded fluorescence in distinct spectral segments, i.e., (i) 430–470 nm and 470–500 nm with filter set 1 and (ii) 430–490 nm and >500 nm with filter set 2 (Figure 3A). This also enabled us to identify mutants that only showed fluorescence changes at certain wavelengths and would have escaped detection, otherwise.

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Characterization of ATP-induced fluorescence changes in the P2X7 head domain recorded at different wavelengths.

(A) Schematic of the voltage clamp fluorometry (VCF)-recording system and summary of filter/dichroic mirror configurations used to detect distinct spectral parts of L-3-(6-acetylnaphthalen-2-ylamino)–2-aminopropanoic acid (ANAP)-fluorescence (sets 1 and 2) and ANAP in combination with tetramethyl-rhodamine-maleimide (TMRM) or R-GECO1.2 (set 3). The corresponding positions A, B, C, and D are shown in the schematic. A second LED (green) was used for additional excitation of TMRM or R-GECO1.2. (B) Close-up of the P2X7 head domain in surface representation indicating the ANAP-substituted amino acid residues P120-Q128 (red). The three subunits are colored in gray, wheat, and light blue. (C) Principle of VCF and representative VCF recordings in response to 0.3 mM ATP (upon second application). Change of fluorescence intensity of a site-specifically introduced environment-sensitive fluorophore can be induced by ligand binding and/or conformational changes. (D) Box plots summarizing results from the indicated ANAP-labeled P2X7Rs at two different emission wavelengths with ΔF/F% representing the maximum fluorescence signal during a 15-s ATP application. Numbers of recordings are given in brackets. (E) Representative VCF recordings in response to 0.3 mM ATP of P2X7(S124*) at three different emission wavelengths and summary of most likely interpretations. Note that fluorescence changes are most likely resulting from multiple effects, and only the dominant effect is stated. Arrows indicate direction of fluorescent changes. (F) Overlay of VCF recordings upon first (colored) and second (gray) ATP applications (0.3 mM) at two different emission wavelengths for P2X7(S124*) (14 oocytes) and P2X7(K127*) (17 oocytes), respectively. Averaged VCF recordings are shown as lines, and standard deviations are plotted as envelopes. Baseline currents (15 s before ATP application) were adjusted for clarity. All recordings were performed in divalent-free buffer, and oocytes were clamped at –30 mV. Original recordings have also been deposited with Dryad and summarized and assigned in Table 1—source data 1.

Figure 3—source data 1 Original recordings, Figure 3C, E and F.: https://cdn.elifesciences.org/articles/82479/elife-82479-fig3-data1-v2.zip
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We first recorded ATP-induced fluorescence changes from P2X7Rs containing ANAP in the head domain, which projects over the ATP-binding site (P120-Q128, Figure 3B). In agreement with the pronounced conformational changes and ligand interactions of this domain during receptor activation (Lörinczi et al., 2012; McCarthy et al., 2019), all mutants except E121* and Q128* showed clear fluorescence signals in all spectral segments (Figure 3C and D, Table 1 and Figure 2—figure supplement 1B). For E121* and Q128* fluorescence changes were only detected with filter set 2, albeit with minimal changes for E121*. Analysis of mutants P120*, E121*, and P123* with filter set 1 was only preliminary (Figure 2—figure supplement 1B) but showed the same trends as signals recorded with filter set 2 (Figure 3C and D). Independently of the wavelength, fluorescence changes were always positive for P120* and G126* and negative for Y122*, P123*, and K127*. These consistent changes over the entire ANAP emission spectrum indicate de-/quenching of ANAP either by the ligand ATP and/or other aa residues (see insert table in Figure 3E). In contrast, S124* (Figure 3E), R125*, and Q128* (Figure 3C and D and Figure 2—figure supplement 1B) showed positive fluorescence changes in most spectral segments but negative changes for wavelengths >500 nm. The opposite directions imply that these changes result, at least partly, from an ANAP emission shift toward shorter wavelengths and suggest that ANAP enters a less polar environment during receptor activation (Figure 3E).

In all head-domain constructs that showed clear kinetics, with the exception of K127*, fluorescence and current changes started simultaneously and showed shorter rising times upon repeated ATP applications, thus recapitulating the characteristic ‘current facilitation’ of the P2X7R (Allsopp and Evans, 2015; Janks et al., 2019; Roger et al., 2008). To confirm that these fluorescence signals indeed tracked current facilitation, we analyzed the effects of three additional mutants that were expected to affect facilitation: (i) a single point mutation in the juxtamembrane region (S23N) that in human P2X7 was shown to eliminate current facilitation (Allsopp and Evans, 2015), (ii) a Cys-Ala mutant (replacement of residues identified to be palmitoylated [McCarthy et al., 2019] in the cysteine-rich region by alanine residues [Ser360, Cys362, Cys371, Cys373, Cys374, and Cys377]), and (iii) a ΔCys-mutant (deletion of the cysteine-rich intracellular region, S360-C377 [McCarthy et al., 2019; Roger et al., 2010]). Contrary to findings in human P2X7 (Allsopp and Evans, 2015), the S23N mutation did not eliminate current facilitation in rat P2X7 (Figure 3—figure supplement 3). As expected (Roger et al., 2010) and in contrast to the wt and most of the analyzed ANAP-containing mutants, current rise times of the ΔCys mutant were not significantly altered between first and second ATP applications, demonstrating that we can indeed identify current facilitation in our setup (Figure 3—figure supplement 3). However, the strongly reduced functional expression of the Cys-Ala or ΔCys mutant with additional ANAP-incorporation sites prevented reliable analysis of current and fluorescence kinetics.

Remarkably, the K127* mutant showed a fluorescence change that was faster than the current increase already upon the first receptor activation (Figure 3F, Figure 3—figure supplement 3). EC₅₀ values for ATP were similar at wt, S124* and K127* mutants (Table 2). We conclude from this, that ANAP in position 127 reports a process that precedes channel opening and is most likely related to ligand binding, whereas ANAP reports gating-associated conformational changes in the other positions.

Table 2

EC₅₀ values for ATP and Hill coefficients (n_H) at wt and L-3-(6-acetylnaphthalen-2-ylamino)–2-aminopropanoic acid-containing P2X7 receptor constructs.

Mutant	EC₅₀ (M)	n_H
Wt	4.202e-005 (3.211e-005–5.704e-005)	1.049 (0.7962–1.380)
F11*	7.802e-005 (6.268e-005–9.893e-005)	1.148 (0.9345–1.410)
S124*	8.316e-005 (6.068e-005–0.0001236)	1.122 (0.8271–1.519)
F11*, S124C	0.0001003 (8.439e-005–0.0001216)	1.290 (1.069–1.571)
K127*	6.511e-005 (4.281e-005–0.0001339)	0.6601 (0.4779–0.8662)
D423*	6.513e-005 (4.729e-005–0.0001057)	1.240 (0.8087–1.821)
A564*	5.159e-005 (3.491e-005–9.976e-005)	0.7810 (0.5364–1.087)

Number in brackets are 95% confidence intervals, n=3–11.

Table 2—source data 1 Original recordings for Table 2; Figure 3—figure supplement 3D; and Figure 5—figure supplement 2A.: https://cdn.elifesciences.org/articles/82479/elife-82479-table2-data1-v2.zip
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Table 2—source data 2 Summarized data for Table 2; Figure 3—figure supplement 3D; and Figure 5—figure supplement 2A.: https://cdn.elifesciences.org/articles/82479/elife-82479-table2-data2-v2.zip
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Detection of TM2 movements in response to receptor activation

The following VCF recordings were performed mainly with filter set 2 (430–490 nm and >500 nm), since this revealed more pronounced signals for most mutants.

To exclude fluorescence changes induced by a direct interaction with ATP and to further investigate P2X7 gating, we next investigated positions 339–341 (Figure 4A), just preceding S342, the major determinant of the channel gate (Pippel et al., 2017). Cysteine substitutions in these positions have previously been shown to be accessible to thiol-reactive dyes only in the open state of the receptor (Pippel et al., 2017). In agreement with a critical role in gating, current recordings from mutants S339*, T340*, and L341* were compromised by 10–20-fold higher leak currents compared to wt receptors or other mutants (see also Figure 4—figure supplement 1). Nevertheless, they showed clear fluorescence changes during receptor activation (Figure 4B), although with higher variability in amplitude and shape between oocytes. While P2X7R mutants S339* and L341* showed positive signals in all spectral ranges, fluorescence changes in T340* were inconsistent at shorter emission wavelengths, but mostly negative below 470 nm, and positive above 470 nm, indicating again that a spectral shift contributed to these signals (Figure 4B and C).

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ATP-induced L-3-(6-acetylnaphthalen-2-ylamino)–2-aminopropanoic acid (ANAP) fluorescence changes in the P2X7 TM2 domain.

(A) Overview and close-up of the three P2X7 subunits (in wheat, gray, and purple) with the TM helices as cartoon representations (in wheat, gray, and green) and the ANAP-substituted residues S339, T340, and L341 (in red). (B) Representative voltage clamp fluorometry (VCF) recordings from the indicated mutants in response to 0.3 mM ATP (upon second application) and summary of results at two different emission wavelengths. Note that recordings from all constructs were compromised by high leak currents. Graphs compare maximal fluorescence signals during first (closed circles) and second (open circles) ATP applications (interval 195 s). Data are represented as mean ± S.E.M. Significance was determined using the two-tailed paired Student’s t-test (*, p<0.05; **, p<0.005). (C) Representative recordings and summary (performed as in B) from P2X7(T340*) with filter set 2. (D) Overlay of VCF recordings from P2X7(T340*) upon first (colored) and second (gray) ATP applications (0.3 mM) at two different emission wavelengths. Averaged VCF recordings from 11 oocytes are shown as lines, and standard deviations are plotted as envelopes. Baseline currents (15 s before ATP application) were adjusted for clarity. All recordings were performed in divalent-free buffer, and oocytes were clamped at –30 mV. Wavelengths passed by the used filter sets are indicated. Original recordings have also been deposited with Dryad and summarized and assigned in Table 1—source data 1.

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Notably, fluorescence signals from P2X7 T340* were also significantly larger during the first ATP application compared to the second (Figure 4B and C), suggesting that the environment of this position changed between both ATP applications. An intriguing explanation could be an involvement of this region in the facilitation process. However, as the T340* mutant displayed no change in fluorescence or current kinetics between ATP applications (Figure 4D and Figure 4—figure supplement 1) the facilitation-associated gating mechanism is likely disturbed by this mutation.

Scanning of the P2X7 intracellular domains for ATP-induced conformational changes

The large intracellular P2X7 C-terminus mediates many of the P2X7R downstream effects (Kopp et al., 2019). While the P2X7 cryo-EM structures revealed the role of the juxtamembrane N- and C-terminal domains in receptor desensitization, their role in downstream signaling and in particular the molecular function of the ballast domain remain completely unclear. Analysis of ANAP fluorescence changes within the cytoplasmic domain was therefore a primary aim of this study. We first introduced ANAP into juxtamembrane regions within the N- and C-termini (Figure 5A) that form the cytoplasmic cap and anchor domains excluding palmitoylated residues (C4, S360, C362, C363, C374, and C377) (McCarthy et al., 2019). Although all receptors with N-terminal ANAP substitutions were formed in full length (Figure 2C), current and fluorescence responses for S6*, V10*, and K17* substitutions remained small and inconsistent even after 6 days of expression. A3*, C5*, W7*, and F11* mutants showed positive fluorescence signals of variable sizes (Figure 5B), and the kinetics of F11* fluorescence correlated with current facilitation (Figure 5C, Figure 3—figure supplement 3). ANAP fluorescence in F11* was not quenched by the nearby Trp (W7), as its removal had no apparent effect (Figure 5—figure supplement 1). Within the juxtamembrane C-terminal regions, ANAP was introduced between TM2 and the anchor domain (N356*, T357*, Y358*, A359*), upstream of β₁₅, which is part of the cytoplasmic cap structure (T361*, R364*, C371*, A378*, R385*, K387*), and upstream of the cytosolic ballast domain (C388* and V392*). Surface expression of functional full-length receptors was observed for all constructs except for K387* and C388*. In contrast to the juxtamembrane N-terminal residues, however, only one of these C-terminal mutants, T361*, showed a fluorescence change, albeit in only ~50% of the recordings (Figure 5D). Interestingly, both F11 and T361 lie within two of at least four possible cholesterol recognition amino acid consensus (CRAC) motifs that have been proposed to be involved in the cholesterol sensitivity of P2X7 channel gating (Robinson et al., 2014).

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L-3-(6-acetylnaphthalen-2-ylamino)–2-aminopropanoic acid (ANAP) incorporation into 41 positions of the cytosolic P2X7 domain identified seven positions that report ATP-induced fluorescence changes.

(A) Surface representation of all three P2X7 subunits (in wheat, gray, and purple) showing location of the juxtamembrane regions and close up (top view) detailing the anchor and cap domains (in yellow and blue, respectively) and ANAP-substituted positions (in red) within a single P2X7 subunit. (B) Representative voltage clamp fluorometry (VCF) recordings and data summary from P2X7R mutants containing ANAP at different positions within the N-terminus. Responses to 0.3 mM ATP were recorded at two different emission wavelengths. Graphs compare maximal fluorescence signals during first (closed circles) and second (open circles) ATP applications (interval 195 s). Data are represented as mean ± S.E.M. (C) Overlay of VCF recordings from P2X7(F11*) upon first (colored) and second (gray) ATP application (0.3 mM) at two different emission wavelengths. Lines represent averaged VCF recordings from 13 oocytes. Standard deviations are plotted as envelopes. Baseline currents (15 s before ATP application) were adjusted for clarity. (D) Representative VCF recordings from the indicated mutants in response to a second application of 0.3 mM ATP and summary of results at the indicated emission wavelengths (performed as in B). Graphs compare maximal fluorescence signals during first (closed circles) and second (open circles) ATP applications (interval 195 s). Data are represented as mean ± S.E.M. All recordings were performed in divalent-free buffer, and oocytes were clamped at –30 mV. (E) Close-up of the cytoplasmic ballast domain from one P2X7 subunit in cartoon and surface representation highlighting a bound GDP (salmon), surrounding α-helices, and residue A564 (red). (F) Surface representation of the cytoplasmic domains of all three P2X7 subunits (in gray, light blue, and wheat) with bound GDP (salmon). Positions in which ATP-induced ANAP fluorescence changes were identified are shown in red. ANAP-substituted positions in which no fluorescence changes were seen (despite surface expression and current responses) are shown in blue. Original recordings have also been deposited with Dryad and summarized and assigned in Table 1—source data 1.

Figure 5—source data 1 Original recordings, Figure 5B and D.: https://cdn.elifesciences.org/articles/82479/elife-82479-fig5-data1-v2.zip
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ANAP introduction in most of the 29 ballast domain positions led to a dominant formation of truncated protein, indicating that this domain does not tolerate substitutions very well and/or that the truncated constructs form stable proteins. Four of these mutants (Q422*, V424*, G538*, I577*) did not form functional receptors at all. For most of the remaining constructs, no specific fluorescence changes could be detected, despite promising surface transport and current responses comparable to wt receptors for at least 12 of them (see Table 1, Figure 2C).

Only in two mutants, A564* and D423*, fluorescence changes could be recorded: A564* showed clearly positive signals, while D423* showed positive signals in only ~40% of the recordings (Figure 5D). Both mutants showed EC₅₀ values similar to wt P2X7 and were not functional in control oocytes injected without ANAP (Figure 5—figure supplement 2), suggesting that the respective truncated proteins (compare Figure 2C) do either not contribute to current responses or only in complex with full-length (ANAP-containing) P2X7 subunits. D423 is located within a loop connecting the β₁₇ and β₁₈ strands and situated on the outer surface of the cytoplasmic ballast, facing away from both the central axis of the receptor and the neighboring subunits (Figure 5F). Notably, mutation of the neighboring positions (Q422*, V424*) resulted in non-functional receptors. A564 is located in the α₁₅ helix at the very end of a cavity formed by the α₁₃, α₁₄, and α₁₆ helices and a short α₉ helix of the neighboring subunit (Figure 5E). This cavity harbors the guanosine nucleotide binding site identified by cryo-EM and liquid chromatography-tandem mass spectrometry analysis, and GDP was found to interact with residues A567 and L569 (McCarthy et al., 2019), both in close proximity to A564. α₁₆, is also part of a proposed lipid interaction or lipopolysaccharide (LPS) binding motif (Denlinger et al., 2001) and α₁₄ at the bottom of the cavity is part of a proposed calcium-dependent calmodulin binding motif (residues I541-S560) (Roger et al., 2010; Roger et al., 2008). To identify possible palmitoylation or CaM-dependent movements of the ballast domain or effects on receptor function, we analyzed the influence of the non-palmitoylated ΔCys- and Cys-Ala mutants (Figure 3—figure supplement 3) as well as a ΔCaM mutant, in which a proposed calmodulin binding site was deleted (Roger et al., 2010) on ANAP fluorescence. While the poor expression of the ΔCys and Cys-Ala mutants in combination with ANAP prevented VCF analysis, combination of the ΔCaM mutation with ANAP (in intracellular positions F11*, D423*, or A564* or in the head domain S124*, K127*) yielded good expression and similar current kinetics and fluorescence changes, as observed before for the single mutants (Figure 5—figure supplement 3). This argues against a major functional effect of the CaM binding site mutation on the current facilitation or on molecular movements, at least in the oocyte-expressed receptor.

Taken together, only two positions, D423 and A564, could be identified within the ballast domain, where ANAP reported environmental changes, suggesting only limited ATP-induced movements in this domain. However, mutant A564* has great potential as a reporter for yet undefined processes that affect GDP binding and/or metabolism.

Parallel recording of ANAP fluorescence with other fluorophores

Based on the above findings, we propose that yet unknown intracellular ligands or protein interactors are required to mediate downstream signaling via the ballast domain. As potential tools to further investigate such molecules and the dynamics of their molecular interplay with the P2X7R, we combined ANAP with other fluorophores and equipped the VCF setup with a second LED for parallel excitation of two different fluorophores within the same protein.

First, we generated a double mutant (F11*/ S124C) suited to investigate the dynamics of P2X7 activation in different parts of the receptor by parallel labeling with the thiol-reactive fluorophore TMRM in the extracellular head domain and with ANAP in the cytoplasmic N-terminus. As seen in Figure 6A, and similar to ANAP in K127*, TMRM in the head-domain showed an instant fluorescent change already upon a first ATP application, whereas the ANAP fluorescence change in F11* was clearly slower. However, both signals coincided upon a second ATP application, further supporting our hypothesis that the so-called current facilitation in P2X7 is due to a change in receptor gating rather than ligand binding. EC₅₀ values for ATP at these mutants and at wt P2X7 were comparable (Figure 3—figure supplement 3 and Table 2).

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Double-labeled P2X7Rs as potential tools to analyze intracellular domain movements, downstream signaling events, and protein interactions.

(A) Scheme of a P2X7 subunit double-labeled with L-3-(6-acetylnaphthalen-2-ylamino)–2-aminopropanoic acid (ANAP) and tetramethyl-rhodamine-maleimide (TMRM) (F11*, S124C) and overlay of fluorescence and current responses to first (colored) and second (gray) ATP applications (0.3 mM) at the indicated emission wavelengths. Lines represent averaged voltage clamp fluorometry (VCF) recordings from five different oocytes and standard deviations are plotted as envelopes. Baseline currents (15 s before ATP application) were adjusted for clarity. (B) Scheme of P2X7(K127*) subunit C-terminally fused to R-GECO1.2 and representative VCF recording in response to 0.3 mM ATP. Recordings were performed in buffer containing 0.5 mM Ca²⁺. (C) Scheme showing the P2X7(Y595*)-CaM-M13-mNeonGreen construct that served as positive control for recordings of FRET between ANAP and mNeonGreen. Ca²⁺ entry through the P2X7R is supposed to induce conformational changes in the CaM-M13-mNeonGreen reporter, which are detected as a FRET signal. A representative VCF recording in response to 0.3 mM ATP is shown. In all recordings, oocytes were clamped at –30 mV.

Figure 6—source data 1 Original recordings, Figure 6A, B, and C.: https://cdn.elifesciences.org/articles/82479/elife-82479-fig6-data1-v2.zip
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Since P2X7 is known to permeate Ca²⁺, an important mediator of intracellular signaling, we also established a protocol to combine VCF recording of ANAP-fluorescence with imaging of P2X7-mediated Ca²⁺ influx by fusing the genetically encoded Ca²⁺-sensor R-GECO1.2 (Wu et al., 2013) C-terminally to the receptor. Combination of P2X7 R-GECO1.2 with the K127* mutant, in which ANAP most likely reports a ligand binding-associated process, showed a clearly delayed onset of the Ca²⁺-dependent R-GECO fluorescence signal, as expected (Figure 6B). A limitation of this protocol was, however, that Ca²⁺ promoted P2X7 desensitization and affected baseline fluorescence, specifically in the ANAP emission spectrum (Figure 6—figure supplement 1). Use of an alternative fluorescent unnatural amino acid (fUAA) would therefore be advantageous.

ANAP has been successfully used as a FRET partner in combination with acceptor transition metals (Gordon et al., 2018), with EGFP (Mitchell et al., 2017), and with YFP to study the apoptosis-regulating Bax-Hsp70 interaction in HeLa cells (Park et al., 2019) and the interaction between BACE1 and KCNQ2/3 in tsA-201 cells (Dai, 2022). Thus, we finally tested whether we could detect FRET signals between ANAP and potential interactors carrying a mNeonGreen-tag. As a proof of concept and based on a CaM-M13-EGFP fusion protein (Mitchell et al., 2017), we generated a positive control (P2X7(Y595*)CaM-M13-mNeonGreen), in which ANAP was introduced into the very C-terminus of a P2X7R that was C-terminally fused to a construct consisting of calmodulin (CaM), CaM-binding myosin light chain kinase (M13), and mNeonGreen (Shaner et al., 2013). Upon Ca²⁺-binding, this construct should move the acceptor protein mNeonGreen in closer proximity to ANAP, which acts as FRET donor.

As expected, ATP-induced Ca²⁺-influx reduced ANAP fluorescence and increased mNeonGreen fluorescence (Figure 6C). The specificity of the signals was confirmed in control experiments (Figure 6—figure supplement 2).

Driven by these results we sought to investigate a potential interaction between the rat P2X7 receptor and CaM (Roger et al., 2010; Roger et al., 2008) and performed experiments with ANAP-labeled P2X7 receptors and mNeonGreen-tagged CaM. However, these recordings revealed no differences to the negative controls, as the CaM-mNeonGreen construct yielded unspecific fluorescence signals (Figure 6—figure supplement 3), possibly due to interaction of soluble mNeonGreen-tagged CaM with the co-injected ANAP.

Since the small FRET signals additionally complicated these analyses, the use of another fUAA with superior photophysical properties such as Acd (Zagotta et al., 2021) might provide a better alternative.

In summary, we identified kinetically different fluorescence changes in the head domain that are most likely associated with ligand binding and gating, respectively, and suggest an involvement of the region around T340 in P2X7 current facilitation. We find, however, only limited ATP-induced movements in the intracellular domains and hypothesize that additional interactions might be required to ‘activate’ the ballast domain. Protocols for parallel recordings of ANAP with TMRM, mNeonGreen, and R-Geco1.2 were established to further analyze such interactions.

Discussion

Optimization of UAA incorporation into P2X7

Site-specific UAA-incorporation represents a powerful method for protein structure-function analysis, and protocols exist for several model systems (Braun et al., 2020; Klippenstein et al., 2018; Leisle et al., 2015; Pless et al., 2015). In X. laevis oocytes, stop codon suppression either by in vitro synthesized UAA-aminoacylated tRNAs or by expression of co-evolved tRNA/aminoacyl-tRNA synthetase pairs has been established. Recently, the semisynthetic ligation of peptide fragments containing the modification using split intervening proteins (inteins) (Sarkar et al., 2021) has also been described (Galleano et al., 2021; Khoo et al., 2020). While chemically aminoacylated tRNA cannot be reloaded after deacylation without a tRNA synthetase (Klippenstein et al., 2018), expression of co-evolved orthogonal tRNA/aminoacyl-tRNA synthetase pairs requires an additional nuclear injection (Kalstrup and Blunck, 2013; Ye et al., 2013). Here, we combined both methods by simultaneously injecting a synthesized suppressor tRNA, cRNA encoding the tRNA synthetase, ANAP, and cRNA encoding the target protein into the cytoplasm. We further enhanced ANAP incorporation by co-injection of cRNA encoding mutated X. laevis eRF1, disfavoring premature translation termination. While mutated eRF1 could potentially interfere with correct translation of endogenous amber-terminated oocyte proteins, we observed no apparent impact on oocyte properties. The presented procedure also improved oocyte quality, expression efficiency, and reproducibility and facilitated optimization of injection ratios. While it does not require equipment for synthesis and purification of UAA-labeled tRNA and is easily applicable in a molecular biology lab, it still depends on a co-evolved tRNA/aminoacyl-tRNA synthetase pair. In combination with UAAs suitable for click chemistry, its flexibility and the choice of fluorophores or functional groups could be greatly expanded (Braun et al., 2020). Here, we could successfully employ the optimized ANAP labeling strategy to explore conformational changes associated with P2X7R activation.

Is P2X7 current facilitation an intrinsic receptor property?

Based on crystal and cryo-EM structures, a molecular mechanism of P2XR gating has been established: ATP-binding to its extracellular inter-subunit binding sites leads to a jaw-like tightening of the head and dorsal fin domains of neighboring subunits around the ATP molecule. This induces an upward movement of β strands in the lower part of the extracellular domain and associated pore opening. Upon prolonged and/or repeated activation, the P2X7R shows a characteristic increase in current amplitude and speed of channel opening, which is generally associated with a shift toward higher ATP sensitivity. Several mechanisms have been proposed to contribute to this so-called current facilitation: modulation of receptor activity by cholesterol (via direct binding to TM domains or cholesterol recognition amino acid consensus [CRAC] motifs) (Karasawa et al., 2017; Murrell-Lagnado, 2017; Robinson et al., 2014), palmitoylation (Di Virgilio et al., 2018; Gonnord et al., 2009; Karasawa et al., 2017), cooperative interactions between intracellular N- and C-termini (Allsopp and Evans, 2015), and calcium-dependent calmodulin binding (Roger et al., 2008). The latter, however, appeared to be specific for rat P2X7 and was not found in the human isoform (Roger et al., 2010). In monocyte-derived human macrophages, current facilitation as well as inflammasome activation, IL-1β release, blebbing, PS flip, and membrane permeabilization were inhibited by phospholipase A2 (PLA2) and Cl^- channel antagonists (Janks et al., 2019), and it was suggested that facilitation represents a downstream effect of P2X7-mediated PLA2 and Cl^- channel activation. Single channel recordings of HEK293 cell-expressed rat P2X7Rs recently revealed an increased open probability as a result of ATP-evoked current facilitation (Dunning et al., 2021). Here, we also observed a faster onset of current signal upon the second ATP application while changes in the amplitude were less obvious. Importantly, for most ANAP-containing P2X7R constructs studied here, fluorescence changes mirrored this behavior, strongly suggesting that it is a receptor-intrinsic property and does not involve currents from downstream-activated channels, such as Ca²⁺-activated Cl^- channels or pannexins (Dunning et al., 2021; Ousingsawat et al., 2015; Pelegrin and Surprenant, 2006; Riedel et al., 2007). Interestingly, the K127* head domain mutant showed faster fluorescence than current changes even upon the first ATP application. Thus, ANAP in this position reports a movement or interaction that precedes channel opening and is most likely related to ligand binding. A similar result was observed for the TMRM-labeled F11*/S124C double mutant, where the onset of TMRM signal upon the first ATP application was faster than the current and ANAP fluorescence change (but coincided upon the second application). In contrast to TMRM, ANAP in position 124 showed fluorescence signals that paralleled current responses, suggesting that different fluorophores can report different processes, possibly due to differences in size and/or sensitivity to the environment. Supporting the idea that these fast fluorescence changes are related to ligand binding, they were only observed in mutants containing fluorophores near the ATP binding site. Fast ligand-induced fluorescence changes already in the non-facilitated P2X7 state imply that ligand binding is unaltered between the first and second activation and consequently, changes in channel gating account for the observed current facilitation.

Fluorescence signals recorded from ANAP in positions near the channel gate (Pippel et al., 2017) could result from different simultaneously occurring effects during channel opening and evidence for both, a shift in ANAP emission toward longer wavelengths (position 340) and dequenching (positions 339 and 341) were observed. Interestingly, ANAP in position 340 revealed significant differences in the fluorescence amplitudes between the first and second ATP application. An intriguing explanation would be that it detects a slowly or non-reversible conformational change after the first ATP application, which could facilitate subsequent gating movements and thereby account for current facilitation. However, T340* was the only construct that did not show a faster current onset upon the second ATP application, possibly because ANAP substitution in this critical position already strongly facilitated gating, as indicated by the large ‘leak’ currents, likely reflecting partial constitutive ligand-independent opening.

Based on the above observations, we propose that the faster activation upon the second ATP application is an intrinsic property of the P2X7R. This conclusion is also in good agreement with the fact that the current facilitation but not downstream signaling events is seen in truncated P2X7 constructs (Kopp et al., 2019; McCarthy et al., 2019). One possibility for a molecular mechanism would be a pre-tensioning of TM2-helices during the first receptor activation that eases channel opening upon a second activation. It is not known, but likely that the cryo-EM structure of the ATP-bound open P2X7R represents the facilitated state. If so, the open-state stabilizing cap domain might not be locked in place in the naïve state but could be formed during the first receptor activation and then stabilized via the cysteine-rich anchor domain. The cap domain may then support the upward transition of TM2 and thereby accelerate current responses. Dynamic cysteine palmitoylation and cholesterol interactions might modulate this process as suggested before (Di Virgilio et al., 2018; Dunning et al., 2021; Karasawa et al., 2017; Robinson et al., 2014). Alternatively, initial receptor activation may change accessibility and/or affinity for a yet unknown allosteric ligand and thereby modulate P2X7 activation. All these suggested mechanisms are not mutually exclusive.

Is the ballast domain affected by ATP-binding/channel opening?

While the functionality of P2X7 as a cation channel is not impaired by lack of the intracellular C-terminus (Becker et al., 2008; Klapperstück et al., 2001; McCarthy et al., 2019), its deletion disrupts a number of P2X7-mediated effects (Kopp et al., 2019), which most likely depend on downstream signaling pathways. A major aim of this study was the identification of C-terminal domains involved in such signaling. Most of the intracellular positions in which ANAP reported relative protein rearrangements were, however, located upstream of the cap domain either within the N-terminus (A3, C5, W7, F11) or right after TM2 (T361). Despite clear surface expression and current responses of at least 12 constructs with ANAP in the cytoplasmic ballast domain, only two of these mutants (D423* and A564*) revealed detectable but small fluorescence changes upon ligand application, suggesting that ATP binding induces only limited structural rearrangement in this domain, and that it is largely uncoupled from the extracellularly initiated conformational changes. Interestingly D423*, which showed only sporadic changes, lies in a short sequence with homology to an α-actinin 2 binding sequence (Kim et al., 2001). Since P2X7 activation induces plasma membrane morphology changes, and interactions with cytoskeletal proteins have been proposed (Gu et al., 2009; Kim et al., 2001; Kopp et al., 2019), an intriguing possibility would be that ANAP in position 423 reports interactions with cytoskeletal components. In A564*, ANAP is located near the GTP/GDP-binding site but showed much smaller signals than in positions near the ATP binding site, arguing against GTP/GDP (un-)binding, in agreement with the cryo-EM structures (McCarthy et al., 2019). However, A564 is also surrounded by other proposed interaction sites, including an LPS binding sequence and a calcium-dependent CaM binding motif (Denlinger et al., 2001; Roger et al., 2010; Roger et al., 2008), which might account for the observed signals.

In summary, we improved ANAP incorporation into Xenopus oocyte-expressed protein and performed an extensive VCF analysis of P2X7R mutants carrying ANAP in 61 positions throughout the receptor. We conclude from our data, that current facilitation is, at least partly, an intrinsic property of the P2X7R and involves an accelerated channel gating rather than ligand binding. In addition, we propose that ligand-induced extracellular and TM domain movements are not significantly translated to the cytosolic ballast domain and that intracellular ligands or interactors are required to ‘activate’ this domain. Protocols for simultaneous recording of ANAP with TMRM, Ca²⁺-dependent R-GECO1.2, or mNeonGreen-labeled FRET partners are presented that might help to validate P2X7 downstream signaling events and analyze their molecular mechanisms and dynamics, once such interactors have been reliably determined.

Materials and methods

Xenopus laevis oocytes

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X. laevis females were obtained from NASCO (Fort Atkinson, WI) and kept at the Core Facility Animal Models (CAM) of the Biomedical Center (BMC) of LMU Munich, Germany (Az:4.3.2–5682/LMU/BMC/CAM) in accordance with the EU Animal Welfare Act. To obtain oocytes, frogs were deeply anesthetized in MS222 and killed by decapitation. Surgically extracted ovary lobes were divided into smaller lobes and dissociated by ~2.5 hr incubation (16°C) with gentle shaking in ND96 solution (96 mM NaCl, 2 mM KCl, 1 mM CaCl₂, 1 mM MgCl₂, 5 mM HEPES, pH 7.4) containing 2 mg/ml collagenase (Nordmark, Uetersen, Germany) and subsequently defolliculated by washing (15 min) with Ca²⁺-free oocyte Ringer solution (90 mM NaCl, 1 mM KCl, 2 mM MgCl₂, 5 mM HEPES). Stage V-VI oocytes were selected and kept in ND96 containing 5 µg/ml gentamicin until further use. In some cases, oocytes were commercially obtained (Ecocyte Bioscience, Dortmund, Germany), or ovaries were provided by Prof. Dr. Luis Pardo (Max Planck Institute for Experimental Medicine, Göttingen, Germany).

cDNA and cloning

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N-terminally His-tagged rat P2X1 cDNA in pNKS2 has been described (Lörinczi et al., 2012). An EGFP-tag was C-terminally added via a GSAGSA-linker sequence by Gibson assembly (Gibson et al., 2009) according to the protocol of the manufacturer (New England Biolabs GmbH, Frankfurt am Main, Germany).

cDNA encoding an N-terminally His-tagged rat P2X7R was subcloned into a pUC19 vector modified for cRNA expression in oocytes (termed pUC19o). pUC19o was generated by insertion (from 5` to 3`) of a synthesized T7 promoter sequence, a Xenopus globin 5’-UTR, and a Kozak sequence (Kozak, 1987) (GeneArt String DNA fragment, Life Technologies / Thermo Fisher Scientific Inc, Regensburg, Germany) and a 27 bp 3’-UTR (Tanguay and Gallie, 1996) followed by a poly A tail (51 adenines) obtained from the pNKS2 vector (Gloor et al., 1995) (for details of the UTRs see Key resource table).

The cDNA sequence of the aminoacyl-tRNA synthetase was obtained from the plasmid pANAP (Addgene #48696) (Chatterjee et al., 2013) and subcloned via Gibson assembly into pUC19o.

The coding sequence of X. laevis eRF1 (NCBI Reference Sequence: NM_001090894.1) with an E55D mutation (GeneArt String DNA fragment, Life Technologies/Thermo Fisher Scientific Inc, Regensburg, Germany) was cloned into pNKS2 via Gibson assembly. For recombinant expression in E. coli, the coding sequence of His-eRF1(E55D) was cloned into a modified pET28a vector via Gibson assembly.

Site-specific mutagenesis was performed with the Q5 Site-Directed Mutagenesis Kit (based on PCR-amplification) according to the manufacturer’s protocol (New England Biolabs GmbH, Frankfurt am Main, Germany). Oligonucleotides were ordered from metabion GmbH (Planegg/Steinkirchen, Germany).

All constructs contained either an ochre (TAA) or opal (TGA) stop codon for normal translational termination to avoid C-terminal ANAP incorporation and read-through and were confirmed by sequencing (Eurofins Genomics, Ebersberg, Germany).

eRF1 protein preparation

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NiCo(DE3) bacteria were transformed with His-eRF1(E55D) in pET28a. 5 ml of a LB-Kanamicin pre-culture (~12 hr) was added to 300 ml ZY-5052 autoinduction media (Studier, 2005) supplemented with 100 µg/ml Kanamycin and grown for 6 hr at 37°C. The temperature was then reduced to 25°C, and bacteria were grown for another 18 hr. After pelleting by centrifugation (6500 g, 20 min) cells were resuspended in 40 ml lysis buffer (50 mM TRIS (tris(hydroxymethyl)aminomethane)-HCl, pH 8.0, 50 mM NaCl, 5 mM MgCl₂, 10% (v/v) glycerol, 0.1% (v/v) Triton X-100, 10 µg/ml DNase I, 100 µg/ml lysozyme), and sonicated (Bandelin Sono plus, TT13 cap, 50% duty cycle, 50% power) for 5 min in an ice bath. The lysate was pelleted at 40,000 × g (1 hr at 4°C). The supernatant was filtered (0.2 µm) and applied onto a Ni-NTA column (HisTrap FF, 5 ml, GE Healthcare Europe GmbH, Freiburg, Germany). Bound protein was washed with 10 column volumes of washing buffer (25 mM TRIS-HCl, pH 7.8, 500 mM NaCl, 20 mM imidazole, 0.25% [v/v] Tween 20, 10% [v/v] glycerol) and eluted with 6 column volumes of elution buffer (25 mM TRIS-HCl, pH 7.8, 500 mM NaCl, 300 mM imidazole, 0.25% [v/v] Tween 20 [v/v], 10% [v/v] glycerol). The eluate was concentrated (Amicon Ultra-15, 10 kDa MWCO, Millipore/Merck KgaA, Darmstadt, Germany), and buffer was exchanged by low-salt buffer (20 mM TRIS, 100 mM NaCl, pH 7.5) for subsequent anion exchange chromatography on a 5 ml Mono-Q column (GE Healthcare Europe GmbH, Freiburg, Germany). Following an elution gradient with high-salt buffer (20 mM TRIS, 1 M NaCl, pH 7.5), protein-containing fractions were pooled, concentrated, and buffer was exchanged (1× PBS with 500 mM NaCl) for size exclusion chromatography on a Superdex 75 Increase (10/300). Purified His-eRF1(E55D) was shock-frozen in 10 µl aliquots and stored at –80°C.

cRNA synthesis and tRNA

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To prepare templates for cRNA synthesis, plasmids were linearized with EcoRI-HF (pNKS2) or NotI-HF (pUC19o) from New England Biolabs GmbH (Frankfurt am Main, Germany) and purified via MinElute Reaction Cleanup columns (Qiagen, Hilden, Germany) according to the manufacturer’s protocol. Alternatively, templates (including the 5’-terminal RNA polymerase promoter site (T7 or SP6) and the 3’-terminal poly A) were amplified by PCR and purified using the NucleoSpin Gel and PCR Clean-up Kit (Macherey-Nagel, Düren, Germany) according to the manufacturer’s protocol.

Capped cRNA was synthesized using the mMESSAGE mMACHINE SP6 or T7 Transcription Kits (Invitrogen/Thermo Fisher Scientific Inc, Schwerte, Germany), precipitated with LiCl, and dissolved in nuclease-free water (1 µg/µl if not stated otherwise).

The amber suppressor tRNA sequence was translated from the plasmid pANAP (Addgene #48696) (Chatterjee et al., 2013), provided with an universal 3’-terminal CCA-sequence (important for tRNA aminoacylation and translation), and chemically synthesized and purified via PAGE and HPLC (biomers.net GmbH, Ulm, Germany).

Oocyte injection and ANAP incorporation

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A Nanoject II injector (Science Products GmbH/Drummond, Hofheim, Germany) was used for nuclear and cytoplasmic injections.

cRNAs encoding cysteine-substituted receptors for TMRM labeling were injected as described (Lörinczi et al., 2012). Two different procedures were used for incorporation of ANAP:

The 2-step injection method was performed according to Kalstrup and Blunck, 2017 using the plasmid pANAP that encodes the co-evolved, orthogonal, and ANAP-specific amber suppressor tRNA/tRNA synthetase pair (Addgene #48696 Chatterjee et al., 2013). 9.2 nl of pANAP (0.1 μg/μl) per oocyte were injected into the nucleus. 1–2 days later, 46 nl of an injection mix containing 0.20–0.25 μg/μl receptor-encoding cRNA (with or without an UAG codon at the site of interest) and 0.2–1.0 mM ANAP (L-ANAP trifluoroacetic salt or L-ANAP methyl ester, both AsisChem Inc, Waltham, MA) were injected into the cytoplasm.

The 1-step injection method was performed as described before (Durner and Nicke, 2022) with addition of mutated X. laevis eRF1 as indicated. An injection master mix comprising 0.25 mM ANAP TFA, 0.25 μg/μl cRNA encoding X. laevis eRF1 E55D, 0.2 μg/μl cRNA encoding the tRNA synthetase, and 0.4 μg/μl tRNA was freshly prepared. Three parts of the injection master mix were added to one part of 1 μg/μl receptor-encoding cRNA (with or without an UAG codon). 50.6 nl per oocyte were injected into the cytoplasm. Uninjected oocytes and oocytes injected with wt receptor cRNA served as negative and positive controls, respectively. Nuclease free water served as a substitute for individual components in control groups.

To optimize fUAA incorporation into X. laevis oocyte-expressed receptors, different procedures, concentrations of substances, and injection time points were compared (Figure 1—figure supplement 1). To optimize the concentrations of an individual component, the ratios and concentrations of the other components, as well as the expression times and receptor cRNA concentrations were kept constant in individual experiments. In cases where oocytes were incubated in membrane-permeable L-ANAP methyl ester, a 2 μM concentration in ND96 buffer (see below) was used.

Injected oocytes were kept in ND96 (96 mM NaCl, 2 mM KCl, 1 mM MgCl₂, 1 mM CaCl₂, 5 mM HEPES, pH 7.4–7.5) supplemented with gentamicin (50 µg/ml) at 16°C for at least 2 days.

Receptor purification and SDS-PAGE

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To evaluate plasma membrane expression of truncated and full-length His-tagged P2X7R mutants, surface-expressed receptors were fluorescently labeled, purified, and analyzed by SDS-PAGE. Three days after injection, 10 oocytes per group were labeled for 30–60 min (in the dark under rotation) in 200 μl 0.003% (m/V) aminoreactive, membrane-impermeant Cy5 Mono NHS Ester (Merck / Sigma-Aldrich, Taufkirchen, Germany, diluted from a 1% [m/V] stock in DMSO) in ND96 (pH 8.5, 4°C) and then washed in ND96. Bright blue-stained damaged oocytes were then discarded, and intact oocytes were homogenized with a 200 μl pipet tip in 10 µl homogenization buffer per oocyte (0.1 M sodium phosphate buffer, pH 8.0, containing 0.4 mM Pefabloc SC and 0.5% n-dodecyl-β-D-maltoside, [both Merck/Sigma-Aldrich, Taufkirchen, Germany]). Membrane proteins were extracted by 10 min incubation on ice and separated from the debris by two centrifugation steps (10 min at 14,000 × g and 4°C). 100 µl of the protein extract were then supplemented with 400 µl of homogenization buffer containing 10 mM imidazole and added to 50 µl Ni²⁺-NTA agarose beads (Qiagen GmbH, Hilden, Germany) preconditioned with washing buffer (0.1 M sodium phosphate buffer [pH 8.0] containing 0.08 mM Pefabloc, 0.1% n-dodecyl-β-D-maltoside, and 25 mM imidazole). After 1 hr incubation under inversion at 4°C in the dark, beads were washed three to four times with 500 µl washing buffer, and His-tagged protein was eluted (≥10 min at RT with occasional flipping to suspend the beads) with 2×50 µl elution buffer (20 mM Tris-HCl, 300 mM imidazole, 10 mM EDTA, and 0.5% n-dodecyl-β-D-maltoside). 32 µl of the eluate were supplemented with 8 µl 5× lithium dodecyl sulfate (LiDS) sample buffer (5% [w/v] LiDS, 0.1% bromphenol blue, 100 mM dithiothreitol, 40% [v/v] glycerol in 0.3 M Tris HCl [pH 6.8]), incubated at 95°C for 10 min, and separated by reducing SDS-PAGE on an 8% gel. Fluorescence-labeled protein was visualized with a Typhoon trio fluorescence scanner (GE Healthcare, Chicago, IL), and relative protein quantities were determined using FIJI (Schindelin et al., 2012). Lanes were selected as regions of interest and transformed into 1D profile plots. Band intensities were then quantified by integrating the area of each peak in the profile plot relative to the baseline of each lane. Data was visualized using GraphPad Prism software (Version 9.3.0, San Diego, CA).

VCF recordings

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Recordings were performed in a custom-made measuring chamber (Figure 3) that is split into an upper and lower compartment, which are individually perfused and connected by a 0.75 mm hole on which the oocyte is placed. The lower compartment has a transparent bottom, and the chamber was mounted on an Axiovert 200 inverted fluorescence microscope (Carl Zeiss Microscopy LLC, Oberkochen, Germany) so that the oocyte was centered above the objective with the animal pole facing down to avoid increased background fluorescence by the lighter vegetal pole. Upper and lower compartments were separately perfused with recording solution and recording or agonist solution, respectively, using a gravity-based perfusion system and a membrane vacuum pump. Solutions in the lower compartment were switched by computer-controlled magnetic valves.

To avoid inhibition by Ca²⁺ or Mg²⁺ and Ca²⁺-mediated downstream effects and to obtain reproducible current responses, recordings were performed in divalent-free buffer (90 mM NaCl, 1 mM KCl, 5 mM HEPES, pH 7.4–7.5) complemented with flufenamic acid and ethylene glycol tetraacetic acid (EGTA) (both 0.1 mM). For measurements with Ca²⁺-containing buffers, EGTA was omitted, and Ca²⁺ (0.2–0.5 mM) was added (in case of P2X7-R-GECO constructs, FRET measurements between ANAP and mNeonGreen and control measurements of ANAP-containing constructs to test for Ca²⁺-specific effects). If not otherwise noted, the agonist solution contained 300 μM ATP and was applied for 15 s in 195 s intervals. Intracellular electrode resistances were below 1.2 MΩ, and recordings were performed at room temperature at a holding potential of –30 mV to keep the current amplitudes reproducible. The solution exchange in the lower chamber is finished in about 1 s (Lörinczi et al., 2012).

To exclude mechanically induced fluorescence changes due to solution switching, all recording protocols started with sequential applications of ATP-free recording solutions from different tubes and magnetic valves. If required, solution speed and oocyte position were readjusted to ensure the absence of mechanical artifacts.

For fluorescence recordings, the microscope was equipped with two LEDs as excitation sources (UV-LED M365LP1 with 365 nm, green LED M565L3 with 565 nm, both Thorlabs GmbH, Bergkirchen, Germany). Since UV excitation in oocytes causes relatively high background fluorescence levels, detectors must feature a wide dynamic range, while maintaining a sufficiently high sensitivity in order to record small fluorescence changes. To this end, two cooled, high-sensitivity MPPC detectors (Hamamatsu Photonics K.K., Japan) were used for simultaneous fluorescence detection at two different spectral segments. For optical filters and dichroic mirrors see Key resource table.

Single-channel fully programmable instrumentation amplifiers with Bessel low-pass filter characteristics (Alligator Technologies, Costa Mesa, CA) were used for signal scaling. To minimize photobleaching, LEDs were pulsed using self-developed high-speed LED drivers with sub-μs rise time. Pulse lengths were set in the ~20 μs range to allow for the fluorescence readout signal chain to settle. Fluorescence signal digitization was synchronized to the excitation pulses using an STM32F407 microcontroller (STMicroelectronics, Geneva, Switzerland). Its timer peripherals were re-triggered by each ADC conversion cycle in order to create an LED illumination pulse that starts shortly before the next ADC conversion cycle. Whenever two excitation wavelengths were used, excitation pulses were staggered in time with the longer wavelength excitation pulse signal being digitized first, preventing bleedthrough of background fluorescence excited by the shorter excitation wavelength to the longer-wavelength detection channel. A water-immersion objective with high numerical aperture and a large working distance (W N-Achroplan 63×/0,9 M27, Carl Zeiss Microscopy LLC, Oberkochen, Germany) was used to maximize the collection of emitted photons and to focus on the oocyte membrane.

Currents were measured with a Turbo Tec-05X amplifier and CellWorks E 5.5.1 software (both npi electronic GmbH, Tamm, Germany) and were used for current and fluorescence recordings and valve control. Current signals were digitized at 400 Hz and downsampled in CellWorks to 200 Hz.

Dose-response analysis

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To determine agonist dose-response curves, ATP was applied for 15 s in 195 s intervals. A reference concentration (ATP_Ref) of 300 μM was applied until stable responses were obtained and was then alternately applied with ATP concentrations ranging from 10 μM to 3 mM (ATP_Test). All responses were normalized to the response of ATP_Ref, and EC₅₀ values were calculated using the four-parameter Hill equation: % Response = Bottom + (Top−Bottom)/(1+10^[(LogEC₅₀−X)*n_H]) with Bottom and Top constrained to 0%, and maximum responses, respectively, X corresponding to the log of agonist concentration, and n_H corresponding to the Hill coefficient.

Data analysis

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Fluorescence and current signals were analyzed and visualized using a Python-based script (for packages used, see Key resource table): Fluorescence signals were denoised using a fifth-order Bessel filter with a low-pass corner frequency of 4 Hz. Maximum amplitudes of ATP-evoked current and fluorescence responses from different receptor constructs were summarized, compared, and visualized using GraphPad Prism software (Version 9.3.0, San Diego, CA). The following inclusion criteria were applied for recordings:

(i) ATP application must evoke a current response >0.1 μA, (ii) leak currents must be stable for the duration of the recording (at least two ATP applications), (iii) repeated ATP applications must elicit reproducible current responses (>0.8 μA), (iv) fluorescence signals must be without mechanical artifacts and clearly distinguishable from fluorescence changes of wt expressing oocytes (see below). 2–3 days after injection, repeated application of 300 μM ATP to wt-expressing oocytes elicited reproducible currents (i.e. first and second current responses differed less than 10% and reached a plateau, at least during the second application), which were taken as a reference. Longer expression times resulted in irregular and irreproducible current responses and less stable oocytes. In case of mutated receptors, longer expression times were often needed to yield current responses comparable to wt P2X7.

We observed a gradual decrease in fluorescence signal for the duration of ligand application in control oocytes expressing wt receptors even in the absence of ANAP. To distinguish ANAP-specific fluorescence signals from these gradual changes, for signal analysis only fluorescence changes upon ATP application were considered that were either positive, or negative but additionally not linear. If fluorescence signals from mutant expressing oocytes were not distinguishable from fluorescence changes observed for wt expressing oocytes no fluorescence change was assumed (0% ΔF/F). Only signals that were recorded in at least three different oocytes were considered for analysis. Additionally, fluorescence changes that were recorded in less than 40% of analyzed oocytes expressing one specific receptor construct or that had averaged ΔF/F values <0.3% were not considered.

Statistical analysis

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Data were either represented as mean ± S.D., as box plots, or as mean ± S.E.M. with the number of recordings given in brackets, and statistical analysis was performed by either two-tailed unpaired Welch’s t-test or two-tailed paired Student’s t-test, as indicated. Values of p<0.05 were defined as statistically significant with *, **, ***, and **** denoting values of p<0.05, 0.005, 0.0005, and 0.0001 or 0.00005, respectively.

Data availability

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Note that original current and fluorescence recordings provided as comma separated value files each contain three columns of values (from left to right): (1) current values, (2) fluorescence signals of longer emission wavelengths, and (3) fluorescence signals of shorter emission wavelengths.

Appendix 1

Appendix 1—key resources table

Reagent type (species) or resource	Designation	Source or reference	Identifiers	Additional information
Recombinant DNA reagent	cDNA Xenopus laevis eRF1(E55D)	NCBI Reference Sequence: NM_001090894.1, Life Technologies / Thermo Fisher Scientific		GeneArt String DNA fragment (cloned into pET28a and pNKS2)
Recombinant DNA reagent	cDNA Rattus norvegicus P2X7			in modified pUC19 (pUC19o)
Recombinant DNA reagent	cDNA Rattus norvegicus P2X1	Lörinczi et al., 2012		in pNKS2
Recombinant DNA reagent	cDNA Rattus norvegicus calmodulin-1	UniProt: PDP29; NCBI Reference Sequence: NM_031969.3		Codon-optimized for Xenopus laevis (Invitrogen / Thermo Fisher Scientific), C-terminally linked to Rattus norvegicus P2X7 via GS-linker (ggatct)
Recombinant DNA reagent	Plasmid: pNKS2	Gloor et al., 1995
Recombinant DNA reagent	Plasmid: pUC19	New England Biolabs GmbH	CAT# N3041S
Recombinant DNA reagent	Plasmid: pANAP	Chatterjee et al., 2013	CAT#48696
Recombinant DNA reagent	EGFP	Yang et al., 1996		the original enhanced GFP, mammalian codon-optimized, C-terminally linked to Rattus norvegicus P2X1 via GSAGSA-linker sequence (ggatctgcaggatctgca)
Recombinant DNA reagent	R-GECO1.2	Wu et al., 2013	Addgene #45494
Recombinant DNA reagent	mNeonGreen	Shaner et al., 2013		Codon-optimized for Xenopus laevis (Invitrogen / Thermo Fisher Scientific)
Recombinant DNA reagent	M13-like peptide from CaM-dependent kinase	Rattus norvegicus myosin light chain kinase, smooth muscle; Uniprot: D3ZFU9		RRKWQKTGNAVRAIGRLSSM; cloned between Rattus norvegicus calmodulin-1 and mNeonGreen with N- and C-terminal GS-linkers (ggcagc and ggatct, respectively)
Sequence-based reagent	Oligonucleotides	metabion GmbH
Sequence-based reagent	Amber suppressor tRNA, synthesized oligonucleotide, sequence derived from pANAP, an universal 3’-terminal CCA-sequence was added: 5'-gcc cgg aug gug gaa ucg gua gac aca agg gau ucu aaa ucc cuc ggc guu cgc gcu gug cgg guu caa guc ccg cuc cgg gua cca –3'	biomers.net GmbH; Chatterjee et al., 2013; Durner and Nicke, 2022
Sequence-based reagent	Sanger sequencing	Eurofins Genomics, https://eurofinsgenomics.eu/
Sequence-based reagent	5’-UTR, GeneArt String DNA fragment (cloned into pUC19 (small letters) before the start codon (italic letters)), gtacccggggatcctctTAATACGACTCACTATAGGCTTGTTCTTTTTGCAGAAGCTCAGAATAAACGCTCAACTTTGGCTCGAGGCCACCatg	Life Technologies / Thermo Fisher Scientific, Kozak, 1987
Sequence-based reagent	3’-UTR, (cloned into pUC19 (small letters) after the stop codon (italic letters)), tgaCCCAAAACAAAAACGGAATATG CAAACAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAAGAATTCTAGAGCGGCCGCagagtcgacctgcagg	pNKS2, Gloor et al., 1995
Peptide, recombinant protein	EcoRI-HF	New England Biolabs GmbH	CAT#R3101S
Peptide, recombinant protein	NotI-HF	New England Biolabs GmbH	CAT#R3189S
Commercial assay or kit	Gibson Assembly Master Mix	New England Biolabs GmbH	CAT#E2611L
Commercial assay or kit	Q5 Site-Directed Mutagenesis Kit	New England Biolabs GmbH	CAT#E0552S
Commercial assay or kit	MinElute Reaction Cleanup Kit	QIAGEN GmbH	CAT#28204
Commercial assay or kit
Commercial assay or kit	Macherey-Nagel NucleoSpin Gel and PCR Clean-up Kit	Fisher Scientific / Thermo Fisher Scientific	CAT# 11992242
Commercial assay or kit	mMESSAGE mMACHINE T7 Transcription Kit	Invitrogen / Thermo Fisher Scientific	CAT# AM1344
Commercial assay or kit	mMESSAGE mMACHINE SP6 Transkription Kit	Invitrogen / Thermo Fisher Scientific	CAT#AM1340
Chemical compound, drug	ATP disodium salt hydrate	Sigma-Aldrich	Cat#A3377
Chemical compound, drug	L-ANAP trifluoroacetic salt	AsisChem Inc.	Cat#ASIS-0014
Chemical compound, drug	L-ANAP methyl ester	AsisChem Inc.	Cat#ASIS-0146
Chemical compound, drug	Collagenase NB 4 G proved grade	Nordmark Pharma GmbH	Cat#S1746502
Chemical compound, drug	Gentamicin sulfate	Roth	CAT#0233.4
Chemical compound, drug	Cy5 Mono NHS Ester	Merck / Sigma-Aldrich	CAT#GEPA15101
Chemical compound, drug	Pefabloc SC	Merck / Sigma Aldrich	CAT#76307
Chemical compound, drug	n-Dodecyl-β-D-Maltoside, ULTROL grade	Merck / Sigma Aldrich	CAT#324355
Chemical compound, drug	Ni-NTA Agarose	QIAGEN GmbH	CAT#1018244
Chemical compound, drug	Flufenamic acid	Merck / Sigma Aldrich	CAT#F9005
Chemical compound, drug	0.5 M EDTA ph 8.0	Thermo Scientific	CAT#R1021
Chemical compound, drug	TMRM	Biomol	CAT#ABD-419
Chemical compound, drug	A 438079 hydrochloride	TOCRIS	CAT#2972
Software, algorithm	CellWorks E 5.5.1	npi electronic, http://cellworks.de/
Software, algorithm	PyMOL	http://www.pymol.org/	RRID:SCR_000305
Software, algorithm	Python Programming Language 3.10.4	http://www.python.org/	RRID:SCR_008394
Software, algorithm	NumPy 1.22.3	http://www.numpy.org	RRID:SCR_008633
Software, algorithm	MatPlotLib 3.5.1	http://matplotlib.sourceforge.net	RRID:SCR_008624
Software, algorithm	SciPy 1.8.0	http://www.scipy.org/	RRID:SCR_008058
Software, algorithm	GraphPad Prism 9.3.0 and 9.5.0	http://www.graphpad.com/	RRID:SCR_002798
Software, algorithm	(Fiji Is Just) ImageJ 2.3.0	Schindelin et al., 2012, http://fiji.sc	RRID:SCR_002285
Other	Turbo Tec-05X Amplifier	npi electronic GmbH	CAT#TEC-05X	VCF-Setup components, electronics
Other	PCI-6221, DAQ, Multifunction I/O Device, 16-Bit	National Instruments	CAT# 779066–01	VCF-Setup components, electronics
Other	Single-channel fully programmable Instrumentation Amplifier Low Pass Filter, USBPGF-S1/L with 8th pole Bessel filter characteristics	Alligator Technologies	CAT#USBPGF-S1/L	VCF-Setup components, electronics
Other	2 x MPPC modules	Hamamatsu Photonics K.K.	CAT#C13366-3050GA	VCF-Setup components, electronics
Other	Power adapter/linear regulator	KNIEL System-Electronic GmbH	Custom-made	VCF-Setup components, electronics
Other	Axiovert 200 inverted fluorescence microscope	Carl Zeiss Microscopy LLC		VCF-Setup components, optics
Other	Objektiv W N-Achroplan 63 x/0,9 M27	Carl Zeiss Microscopy LLC	CAT#420987-9900-000	VCF-Setup components, optics
Other	M565L3, mounted LED at 565 nm	Thorlabs GmbH	CAT#M565L3	VCF-Setup components, optics
Other	M365LP1, Mid Power Mounted LED at 365 nm	Thorlabs GmbH	CAT#M365LP1	VCF-Setup components, optics
Other	2× lenses for LED collimation	Thorlabs GmbH	CAT#ACL2520U-A	VCF-Setup components, optics
Other	ET555/20×, 25 mm Dia Mounted, Single Bandpass Filter (for excitation)	Chroma Technology GmbH	CAT# IN026697	VCF-Setup components, optics
Other	ET365/20×, 25 mm Dia Mounted, Single Bandpass Filter (for excitation)	Chroma Technology GmbH	CAT# IN053211	VCF-Setup components, optics
Other	T387lp, 25.5×36×1 mm, Longpass Dichroic Beamsplitter	Chroma Technology GmbH	CAT# IN040921	VCF-Setup components, optics
Other	79003bs, Multi Dichroic Beamsplitter	Chroma Technology GmbH	CAT# CS294227	VCF-Setup components, optics
Other	59002bs, Multi Dichroic Beamsplitter	Chroma Technology GmbH	CAT# IN040206	VCF-Setup components, optics
Other	T425lpxr,25.5×36×1 mm, Longpass Dichroic Beamsplitter	Chroma Technology GmbH	CAT# IN025246	VCF-Setup components, optics
Other	Relay lense	Thorlabs GmbH	CAT#AC254-060-A	VCF-Setup components, optics
Other	DMLP550R, Longpass Dichroic Beamsplitter	Thorlabs GmbH	CAT#DMLP550R	VCF-Setup components, optics
Other	T470lpxr, Longpass Dichroic Beamsplitter	Chroma Technology GmbH	CAT# IN030502	VCF-Setup components, optics
Other	T495lpxr, 25.5×36×1 mm, Longpass Dichroic Beamsplitter	Chroma Technology GmbH	CAT# IN005752	VCF-Setup components, optics
Other	ET490/40×, 25 mm Dia Mounted (for emission)	Chroma Technology GmbH	CAT# IN039532	VCF-Setup components, optics
Other	ET610/75 m, 25 mm Dia Mounted (for emission)	Chroma Technology GmbH	CAT# IN036520	VCF-Setup components, optics
Other	ET455/50 m (for emission)	Chroma Technology GmbH	CAT# IN067607	VCF-Setup components, optics
Other	ET500lp, 25 mm Dia Mounted (for emission)	Chroma Technology GmbH	CAT# IN006640	VCF-Setup components, optics
Other	MF460-60 (for emission)	Thorlabs GmbH	CAT#MF460-60	VCF-Setup components, optics
Other	2× lenses for focusing on MPPC	Thorlabs GmbH	CAT#LB1761-A	VCF-Setup components, optics

Data availability

All data generated or analyzed during this study are included in the manuscript and supporting files. Original recordings or scans from SDS-PAGE gels are provided as source data with the respective figures. Original recordings of Table 1 are deposited with Dryad (for data assignment see Table 1-source data 1). This paper does not report original code.

The following data sets were generated

(2023) Dryad Digital Repository
Table 1-source data 1 (original recordings).
https://doi.org/10.5061/dryad.p8cz8w9tb

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Decision letter

Stephan A Pless

Reviewing Editor; University of Copenhagen, Denmark
Richard W Aldrich

Senior Editor; The University of Texas at Austin, United States
Gucan Dai

Reviewer; Saint Louis University, United States

Our editorial process produces two outputs: (i) public reviews designed to be posted alongside the preprint for the benefit of readers; (ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "Voltage clamp fluorometry analysis of the P2X7 receptor suggests a limited conformational interplay between extracellular ATP binding and the intracellular ballast domain" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Richard Aldrich as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Gucan Dai (Reviewer #3).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

The reviewers appreciated the effort needed to generate this impressive data set. Although there was enthusiasm for the overall approach and the reviewers agreed on the potential of the study, it was felt that additional mechanistic insight on facilitation and impact of intracellular factors would be required before the work would be of sufficient interest. Therefore we ask you to carefully address the following essential revisions:

(1) Examination of fluorescence changes in the ballast domain (e.g. A564*) in response to manipulation of at least one or two relevant cytoplasmic factors (e.g. GTP/GDP, Zn2+, palmitoylation (e.g. mutations) or Ca²⁺/CaM). This is crucial to provide mechanistic insight on the functional role of the ballast domain.

(2) Careful validation that the fluorescence changes from e.g. S124*, F11*, S127C do in fact track facilitation. This could be achieved by studying fluorescence signals of mutants that alter or eliminate facilitation (if those mutants exist) or by examining the effect of experimental manipulations of cytoplasmic factors on fluorescence (and current) facilitation (e.g. removal of extracellular Ca²⁺ or similar). This is particularly interesting for the intracellular N-terminus, which cannot be labeled with cysteine-reactive fluorophores.

(3) A more thorough characterization of key mutants in terms of ATP sensitivity of current and fluorescence, facilitation and activation/deactivation kinetics. This is required for a more quantitive analysis and has the potential to yield additional insight on the mechanism of receptor activation.

Reviewer #1 (Recommendations for the authors):

I would have expected the authors to functionally characterize the mutants on which they focus in the paper, for instance by measuring the ATP EC50, to have an idea of whether ANAP incorporation at the different sites perturbs receptor function.

When studying fluorescence changes at the level of the head domain (p7), the authors write that "fluorescence and current changes started simultaneously and showed shorter rising times upon repeated ATP applications"; "K127* showed a fluorescence change that was faster than the current increase" but no quantification of the fluorescence and current kinetics is given. It would be nice to have such quantification. It would be also nice to have a comparison of the fluorescence and current ATP dose-response curves for mutants showing sufficiently large fluorescence changes. This would further support the fact that ANAP at the level of the head domain tracks gating.

ANAP incorporation at TM2 positions seems to induce constitutive currents in P2X7Rs. Is there a channel blocker specific for P2X7Rs that would allow to give an estimation of constitutive activity of these mutants?

Page 9, it is written that "the kinetics of F11* fluorescence correlated with current facilitation". As before, this statement solely relies on the qualitative observation of current and fluorescent traces. Quantitative measurements of fluorescence and current kinetics during the first and second application of ATP is necessary to substantiate that claim. Here again, a dose-response curve of current and fluorescence for ATP of P2X7-F11* would be a plus to determine the gating step(s) tracked by ANAP at this position.

Attributing the lack of ATP-induced fluorescence changes observed in mutants with ANAP incorporated in the ballast domain to a lack of functional coupling between the extracellular and the intracellular domains is a strong assumption. The issue with ANAP incorporation in the C-terminus is the membrane expression of truncated proteins that are functional. Since the authors showed that these truncated proteins exists, if they are functional, they contribute to the measured current. The proportion of ANAP-labeled proteins could thus be too low to yield fluorescence changes despite large currents. To study the contribution of truncated proteins, the authors should characterize these truncated proteins by expressing just the mRNA with the amber stop codon in Xenopus oocytes, investigate potential ATP-induced currents carried by these truncated receptors and measure ATP dose-response curves. Comparing ATP dose-response curves of ANAP-mutants to purely truncated proteins should allow estimating the contribution of truncated proteins to the general current.

It seems that calcium induces fluctuations of ANAP fluorescence. Does calcium have any direct effect on ANAP fluorescence?

It seems that fluorescence changes at some positions of the extracellular head domain and intracellular N-terminus track the process of facilitation. Tracking of this process is probably the most interesting finding of this paper, since the mechanism of facilitation is not known. In the paper, facilitation is measured as the increase of channel opening kinetics upon multiple applications of ATP. However, the first application of ATP clearly shows a small, fast component and a bigger, slow component (see, for instance, Figure 3F, 5C, 6A). Could it be that the first component reflects initial gating and the second one the slow facilitation process that occurs upon prolonged ATP application? Could the fast kinetics of K217* fluorescence match the fast component of current, indicating that K217* tracks the initial ATP opening step but not current facilitation? Similarly, on Figure 6 it looks like TMRM kinetics match first the fast current component, while F11* fluorescence matches the kinetics of current facilitation. Could the authors comment on it?

Reviewer #2 (Recommendations for the authors):

To strengthen the points of weakness above and to elevate the scientific merits of the paper, I have some comments which require attention.

(1A) I am curious to know the P2X7 structural differences by VCF between just before the 1st application and just before the 2nd application of ATP, to clarify the structural background of "memory" of ATP application.

(1B) Will there be any difference in (1A), in the presence and absence of extracellular Ca²⁺, i.e. Ca²⁺ influx via P2X7.

(2A) Facilitation of F change was detected e.g. in S124*, F11*, S127C. Possible mechanisms of facilitation are discussed including mediators such as GDP, cholesterol, palmitoylation, Ca²⁺/CaM (Lines 335-342), However, there are no experimental results to support or deny them. It is desired to examine the effect of experimental manipulations of these factors on the F(and I) facilitation (e.g. removal of extracellular Ca²⁺).

(2B) Line 273: "the so-called current facilitation in P2X7 is due to a change in receptor gating rather than ligand binding", Line 366: "Ligand binding is unaltered between the 1st and 2nd activation and consequently, changes in channel gating account for the observed current facilitation". I wonder if this is correct, as F facilitation is observed in S124* which is close to the ATP binding site (Figure 3F).

(2C) F facilitation was detected not in S124C-TMRM (Figure 6A), not in K127* (Figure 3F), but in S124* (Figure 3F). I cannot fully understand the ground of differences.

(3) As to the function of the ballast domain, it is described "intracellular ligand or interactors are required to "activate" this domain" (line 421). A possible involvement of the ballast domain to the functional modulation of P2X7 by the binding of various factors such as GTP/GDP, LPS and Ca²⁺/CaM are discussed (lines 407-415). The authors showed limited structural rearrangements of the ballast domain upon ATP application, but there is no evidence by VCF to prove activation by intracellular ligand. I would like to see data of F change of such as A564* in the ballast domain in response to manipulation of intracellular factors.

(4) Figure 4C: Does the F facilitation in T340* truly reflect "facilitation", in spite of the absence of current facilitation (as described in line 376)? The authors explained that a lack of faster current in the 2nd application is due to "strongly facilitated gating" (line 377). If that is the case, why there is F facilitation?

(5) Do the F change of R-Geco (Figure 6B) and FRET change by mNeonGreen (Figure 6C) disappear in the absence of extracellular Ca²⁺?

Reviewer #3 (Recommendations for the authors):

1. Please cite this paper when talking about eRF-E55D as well, https://doi.org/10.7554/eLife.37248.

2. The 3'-terminal CCA-sequence is already in the pANAP plasmid. More elaboration on how it augmented the translation is needed to make it clear to readers.

3. Additional FRET experiments might be out of the scope of this paper, but please add more discussion and references regarding this possibility in the paper.

4. One area that could be improved on is the uncoupled "ballast" domain. How might GTP/GDP and dinuclear Zn2+-binding rearrange this domain? Can pharmacological manipulations be used to change intracellular GTP/GDP or Zn2+ levels to see how Anap labeled in this domain changes its fluorescence differently? Could the palmitoylation of the anchor domain play a role for uncoupling the "ballast' domain and for the current facilitation?

5. Line 634, should be "artifacts"?

https://doi.org/10.7554/eLife.82479.sa1

Author response

Essential revisions:

The reviewers appreciated the effort needed to generate this impressive data set. Although there was enthusiasm for the overall approach and the reviewers agreed on the potential of the study, it was felt that additional mechanistic insight on facilitation and impact of intracellular factors would be required before the work would be of sufficient interest. Therefore we ask you to carefully address the following essential revisions:

(1) Examination of fluorescence changes in the ballast domain (e.g. A564*) in response to manipulation of at least one or two relevant cytoplasmic factors (e.g. GTP/GDP, Zn2+, palmitoylation (e.g. mutations) or Ca²⁺/CaM). This is crucial to provide mechanistic insight on the functional role of the ballast domain.

We agree that more insight in the role of the ballast domain would be highly informative. However, the relevance of the cytoplasmic factors such as Zn²⁺ and GTP has not been defined so far and no functional readouts for mutations in the GTP or Zn²⁺ binding sites are known, which hampers their investigation. At least in the Zn²⁺ binding site, we expected mutations to be not tolerated (Gonnord et al., 2009), likely because of a structural function in receptor folding. Also, our set-up and experimental conditions do not allow a direct comparison of effects before and after intracellular modulation (e.g. by Zn²⁺ or GDP/GTP).

We therefore concentrated on possible effects of (de-)palmitoylation and a previously described interaction with calmodulin and generated and analyzed the following three additional mutants:

(i) Cys-Ala P2X7 (replacement of six palmitoylated residues Ser360 and Cys362, 371, 373, 374, 377, McCarthy et al., 2019) in the cysteine-rich region by alanine residues,

(ii) DCys P2X7 (deletion of the cysteine-rich intracellular region (S360-C377)), and

(iii) DCaM P2X7 (disruption of a proposed calmodulin (CaM) binding site by three point mutations I541T, S552C, V559G Roger et al., 2010).

According to (McCarthy et al., 2019) and (Roger et al., 2010), DCys- and Cys-Ala mutants were expected to be not palmitoylated and to show no facilitation. In agreement with this hypothesis, we indeed did not see facilitation in the DCys mutant while expression of the Cys-Ala mutant was too low for analysis (Figure 3—figure supplement 3, compare also point 2). Expression of these mutants, was further reduced by the additional mutations for ANAP incorporation (F11*, S124*, K127*, D423*, or A564*), making a reliable detection of the corresponding fluorescence changes impossible.

The CaM-binding site in rat P2X7 was previously shown to affect current facilitation (Roger et al., 2010). Here, the DCaM mutant showed facilitation comparable to wt P2X7. Likewise, combination with ANAP in intracellular positions or the head domain (F11*, D423*, A564* and S124*, K127*, respectively) revealed similar fluorescence changes as observed for the single mutants with intact CaM binding sites. This argues against a major functional effect of the CaM binding site mutation on current kinetics or molecular movements in the oocyte-expressed receptor. These data are now included in Figure 5—figure supplement 3 and mentioned in line 286ff.

As an additional approach to address a potential interaction of the P2X7 receptor with CaM, we performed FRET experiments with mNeonGreen-tagged CaM and ANAP-labeled P2X7 (ANAP introduced in A3, Q455, L523, or L527 and preliminary experiments with R125, T340, R364, E406, V517, L569, and Y595). However, these recordings revealed no differences to the negative control (wt P2X7 with co-injected ANAP and mNeonGreen-tagged CaM) and were therefore discontinued.

These additional data are now added as Figure 6—figure supplement 3 and referred to in the text (lines 343ff).

Although these experiments cannot exclude an interaction with CaM, they do not support an important role of CaM in the observed current facilitation.

(2) Careful validation that the fluorescence changes from e.g. S124*, F11*, S127C do in fact track facilitation. This could be achieved by studying fluorescence signals of mutants that alter or eliminate facilitation (if those mutants exist) or by examining the effect of experimental manipulations of cytoplasmic factors on fluorescence (and current) facilitation (e.g. removal of extracellular Ca²⁺ or similar). This is particularly interesting for the intracellular N-terminus, which cannot be labeled with cysteine-reactive fluorophores.

Deletion of the cysteine-rich region was previously shown to eliminate current facilitation (Roger et al., 2010). In agreement, current rise times (compare also point 3 below) of the first and second ATP applications to the DCys mutant were unaltered in our experiments. In contrast, current rise times clearly accelerated in the wt, S124*, and F11* mutations, demonstrating that we can indeed identify current facilitation in our set-up. However, the DCys and Cys-Ala mutants showed already decreased currents and their combination with sites for ANAP incorporation (F11*, S124*) resulted in further reduced functional expression, which prevented analysis of the corresponding fluorescence changes.

An S127C mutant was not prepared or mentioned in the text and we assume that the reviewer referred to K127*, which showed facilitation only for the current but not for the fluorescence. For DCys/K127*, robust fluorescence changes were only detected in one out of 12 oocytes. In this case, the fluorescence changes were virtually identical upon first and second ATP application, as expected.

As an alternative approach, we also combined the three ANAP-substituted mutants (S124*, K127*, F11*) with an S23N mutant that has been shown to alter the kinetics of receptor activation in human P2X7 (Allsopp and Evans, 2015). However, this mutation did not induce an effect on rat P2X7 kinetics.

The analyses confirming facilitation of current rise times in the S124*, F11* mutations and elimination of current facilitation by the DCys mutation are now included in Figure 3—figure supplement 3 and in lines 190ff in the text.

Extracellular Ca²⁺ was generally omitted in the presented VCF recordings since it interfered with ANAP fluorescence measurements (compare line 324 and material and methods, line 653ff). Ca²⁺-containing buffer affected overall baseline fluorescence in VCF-experiments independent of the presence or absence of ANAP and additionally promoted receptor desensitization (Figure 6—figure supplement 1), making these conditions unsuitable for reliable detection and analysis of receptor responses.

We now added photometric data, demonstrating also a direct effect of Ca²⁺ on ANAP fluorescence (Figure 6—figure supplement 1), albeit only at very high concentrations.

(3) A more thorough characterization of key mutants in terms of ATP sensitivity of current and fluorescence, facilitation and activation/deactivation kinetics. This is required for a more quantitive analysis and has the potential to yield additional insight on the mechanism of receptor activation.

We generated ATP dose response curves for the current responses of wt P2X7 receptor, the key mutants F11*, S124*, K127*, D423*, A564*, and the TMRM-labeled F11*/S124C double mutant and show that their ATP sensitivities differed less than 3fold from that of the wt P2X7.

Unfortunately, dose response analysis of fluorescence responses could not be performed due to ANAP bleaching during the long recording protocol (about 15-40 min) and the high variability of background fluorescence, which made normalization of responses from different oocytes impossible.

For a quantitative analysis of current facilitation/activation kinetics, we determined the 10-50% rise times of current and fluorescent signal increases upon first and second ATP application for wt P2X7, the DCys mutant, the DCaM mutant, as well as mutants F11*, S124*, K127*, T340*, and the TMRM-labeled F11*/S124C double mutant. Kinetics of channel deactivation were not analyzed as they were strongly delayed, most likely due to turbulences in the measuring chamber that resulted in a slow wash-out of ATP.

These data are now provided in Figure 3—figure supplement 3, Figure 4—figure supplement 1, Figure 5—figure supplement 2 and 3, and Table 2. For corresponding text see specific reviewer questions below.

Reviewer #1 (Recommendations for the authors):

I would have expected the authors to functionally characterize the mutants on which they focus in the paper, for instance by measuring the ATP EC50, to have an idea of whether ANAP incorporation at the different sites perturbs receptor function.

ATP EC₅₀ values of current responses have now been added for the wt P2X7 receptor and key mutants F11*, S124*, K127*, D423*, A564* as well as the F11*/S124C double mutant. EC₅₀ values at the mutants differed less than 3-fold from that at the wt. These data have been included in Table 2, Figure 3—figure supplement 3, and Figure 5—figure supplement 2 and were referred to in lines 208, 271, 315.

When studying fluorescence changes at the level of the head domain (p7), the authors write that "fluorescence and current changes started simultaneously and showed shorter rising times upon repeated ATP applications"; "K127* showed a fluorescence change that was faster than the current increase" but no quantification of the fluorescence and current kinetics is given. It would be nice to have such quantification. It would be also nice to have a comparison of the fluorescence and current ATP dose-response curves for mutants showing sufficiently large fluorescence changes. This would further support the fact that ANAP at the level of the head domain tracks gating.

ATP dose response curves for currents have been added (see above). Generation of dose response curves for fluorescence was hampered by bleaching and varying background fluorescence of individual oocytes (see also above and Essential Revision, point 3).

Quantification of current and fluorescence kinetics for the key mutants F11*, S124*, K127* and the TMRM-labeled F11*/S124C double mutant has now been performed and the respective data have been included in Figure 3—figure supplement 3 (see also Essential Revision, point 2).

ANAP incorporation at TM2 positions seems to induce constitutive currents in P2X7Rs. Is there a channel blocker specific for P2X7Rs that would allow to give an estimation of constitutive activity of these mutants?

To our knowledge, there is no channel blocker of P2X7 available. However, the P2X7 antagonist A438079 (Allsopp et al., 2018) was able to decrease the leak currents of the T340* and L341* mutants. These data are now included as Figure 4—figure supplement 1 (referred to in line 223).

Page 9, it is written that "the kinetics of F11* fluorescence correlated with current facilitation". As before, this statement solely relies on the qualitative observation of current and fluorescent traces. Quantitative measurements of fluorescence and current kinetics during the first and second application of ATP is necessary to substantiate that claim. Here again, a dose-response curve of current and fluorescence for ATP of P2X7-F11* would be a plus to determine the gating step(s) tracked by ANAP at this position.

These data have now been included (see above and Essential Revision, point 2).

Attributing the lack of ATP-induced fluorescence changes observed in mutants with ANAP incorporated in the ballast domain to a lack of functional coupling between the extracellular and the intracellular domains is a strong assumption. The issue with ANAP incorporation in the C-terminus is the membrane expression of truncated proteins that are functional. Since the authors showed that these truncated proteins exists, if they are functional, they contribute to the measured current. The proportion of ANAP-labeled proteins could thus be too low to yield fluorescence changes despite large currents. To study the contribution of truncated proteins, the authors should characterize these truncated proteins by expressing just the mRNA with the amber stop codon in Xenopus oocytes, investigate potential ATP-induced currents carried by these truncated receptors and measure ATP dose-response curves. Comparing ATP dose-response curves of ANAP-mutants to purely truncated proteins should allow estimating the contribution of truncated proteins to the general current.

Thank you for the helpful suggestion, these control experiments have now been performed for the D423* or A564* mutants and revealed less than 5% of the respective control current responses (RNA co-injected with all essential components) if the respective RNAs were co-injected only with water.

Thus, these truncated subunits do not or very inefficiently form functional homotrimers and their contribution to the current can be largely excluded. However, they might form functional heterotrimers with ANAP-labeled full-length subunits and as such, could still contribute to the fluorescence signal. Nevertheless, the fact that the EC₅₀ values at the ANAP-labeled constructs were similar to those at the wt P2X7, argues against a major effect of the truncated subunits.

These data have now been included as Figure 5—figure supplement 2.

It seems that calcium induces fluctuations of ANAP fluorescence. Does calcium have any direct effect on ANAP fluorescence?

To test this possibility, we measured the emission spectra of ANAP-containing recording solution in the absence and presence of 10 μM, 100 μM, 1 mM, 10 mM, 100 mM, or 1 M Ca²⁺ and found that Ca²⁺ increases ANAP fluorescence, but only at concentrations above 100 mM. We included these photometric data in Figure 6—figure supplement 1.

It needs to be considered, however, that the Ca²⁺-induced fluctuations in VCF-measurements occurred also in the absence of ANAP (Figure 6—figure supplement 1).

It seems that fluorescence changes at some positions of the extracellular head domain and intracellular N-terminus track the process of facilitation. Tracking of this process is probably the most interesting finding of this paper, since the mechanism of facilitation is not known. In the paper, facilitation is measured as the increase of channel opening kinetics upon multiple applications of ATP. However, the first application of ATP clearly shows a small, fast component and a bigger, slow component (see, for instance, Figure 3F, 5C, 6A). Could it be that the first component reflects initial gating and the second one the slow facilitation process that occurs upon prolonged ATP application? Could the fast kinetics of K217* fluorescence match the fast component of current, indicating that K217* tracks the initial ATP opening step but not current facilitation? Similarly, on Figure 6 it looks like TMRM kinetics match first the fast current component, while F11* fluorescence matches the kinetics of current facilitation. Could the authors comment on it?

Indeed, current changes appear to be biphasic and we cannot exclude that the fast current component from K127* and TMRM-labeled F11*/S124C P2X7 match initial fast gating that occurs before the onset of facilitation and is reported by ANAP in position 127 or TMRM in position 124. However, the initial fast current component increases substantially between first and second ATP-application. This is not mirrored by the fluorescence signals, which are virtually identical between first and second ATP-applications. Therefore, we think that the fluorescence for this mutant reports a different process. Also, taking into account the proximity of the fluorophores to the orthosteric binding-site we consider it more likely, that these mutants report a process related to ligand binding. Compare also new Figure 3—figure supplement 3.

Reviewer #2 (Recommendations for the authors):

To strengthen the points of weakness above and to elevate the scientific merits of the paper, I have some comments which require attention.

(1A) I am curious to know the P2X7 structural differences by VCF between just before the 1st application and just before the 2nd application of ATP, to clarify the structural background of "memory" of ATP application.

Comparing the structures before first and second ATP application is very difficult with an indirect method like VCF, which provides mainly information about dynamic processes. Determination of different receptor states with this method would require clearly distinct fluorescence signals upon first and second activation, detailed structural information, and most likely numerous additional mutants to deduce and confirm specific conformational changes. Based on our observations, we generally suggest a potential transition into a pre-activated state (line 446) but do not want to elaborate on this to avoid overinterpretation.

(1B) Will there be any difference in (1A), in the presence and absence of extracellular Ca²⁺, i.e. Ca²⁺ influx via P2X7.

Experiments in presence of extracellular Ca²⁺ (1 mM) were performed to test for Ca²⁺-specific effects (Figure 6—figure supplement 1) but revealed large unspecific signals (even in the absence of ANAP) in the spectral range of ANAP emission and thus hampered analysis. Recordings in 0.5 mM Ca²⁺ from oocytes expressing F11* and S124* did not reveal obvious changes in the pattern of ANAP fluorescence signals compared to measurements in divalent-free recording solution.

(2A) Facilitation of F change was detected e.g. in S124*, F11*, S127C. Possible mechanisms of facilitation are discussed including mediators such as GDP, cholesterol, palmitoylation, Ca²⁺/CaM (Lines 335-342), However, there are no experimental results to support or deny them. It is desired to examine the effect of experimental manipulations of these factors on the F(and I) facilitation (e.g. removal of extracellular Ca²⁺).

We performed additional experiments to investigate effects of palmitoylation (using two mutants in which the palmitoylation sites were removed, Figure 3—figure supplement 3) and a CaM interaction using a mutant with mutated CaM-binding motif and investigating FRET between ANAP-labeled P2X7 and mNeonGreen-tagged CaM (Figure 5—figure supplement 3 and Figure 6—figure supplement 3, respectively). These data confirmed that deletion of a cysteine-rich intracellular region eliminates current facilitation (Roger et al., 2010) and that some of our mutants indeed track facilitation. However, mutation of the CaM binding site and FRET experiments did not support an effect of calmodulin or were inconclusive. Please see Essential Revisions points 1and 2 for details.

We would also like to point out that the changes observed in our study are compatible with a facilitation process that is an intrinsic property of the channel and does not require intracellular factors.

(2B) Line 273: "the so-called current facilitation in P2X7 is due to a change in receptor gating rather than ligand binding", Line 366: "Ligand binding is unaltered between the 1st and 2nd activation and consequently, changes in channel gating account for the observed current facilitation". I wonder if this is correct, as F facilitation is observed in S124* which is close to the ATP binding site (Figure 3F).

As described, the fluorescence changes can report both ligand interactions and protein movements as well as combinations of both. The specific effects depend not only on the orientation and position of the mutated residue but also the fluorophore used (e.g. different kinetics of TMRM and ANAP in position 124, compare also line 420ff). The fact that some mutants, like TMRM-labeled double mutant F11*/S124C and K127* detect fast processes (likely associated with direct ligand interactions or movements during ligand binding), does not exclude the possibility that fluorophores in neighboring positions (like ANAP-labeled S124*) detect relative domain movements within the receptor (likely resulting from a quenching/dequenching by aromatic residues or changes in the polarity of the environment) that contribute to the opening mechanism.

(2C) F facilitation was detected not in S124C-TMRM (Figure 6A), not in K127* (Figure 3F), but in S124* (Figure 3F). I cannot fully understand the ground of differences.

ANAP can detect localized changes in its environment in a distance of about 5 Å. Therefore, the immediate environment can be completely different for S124* and K127* and ANAP can even report different changes in neighbouring positions. In case of ANAP-labelled S124* and TMRM-labeled S124C, the very differently sized fluorophores most likely sense different environments and therefore also report different interactions/effects (quenching or polarity).

(3) As to the function of the ballast domain, it is described "intracellular ligand or interactors are required to "activate" this domain" (line 421). A possible involvement of the ballast domain to the functional modulation of P2X7 by the binding of various factors such as GTP/GDP, LPS and Ca²⁺/CaM are discussed (lines 407-415). The authors showed limited structural rearrangements of the ballast domain upon ATP application, but there is no evidence by VCF to prove activation by intracellular ligand. I would like to see data of F change of such as A564* in the ballast domain in response to manipulation of intracellular factors.

In contrast to patch-clamp experiments, direct manipulation of intracellular factors during the recording is difficult if not impossible in TEVC experiments. As an alternative, we generated mutations (as suggested) in the palmitoylation site and the CaM binding site and performed FRET experiments with co-expressed ANAP-labeled P2X7 and mNeonGreen-tagged CaM. We further did experiments in different concentrations of extracellular Ca²⁺. For details, please see Essential revision points 1 and 2. These data are now included in Figure 3—figure supplement 3, Figure, Figure 5—figure supplement 3, Figure 6—figure supplement 1, and Figure 6—figure supplement 3.

(4) Figure 4C: Does the F facilitation in T340* truly reflect "facilitation", in spite of the absence of current facilitation (as described in line 376)? The authors explained that a lack of faster current in the 2nd application is due to "strongly facilitated gating" (line 377). If that is the case, why there is F facilitation?

The beauty of VCF is the ability to differentiate between electrophysiologically defined receptor states (open and closed) and associated domain movements. It is well possible, that due to a "leaky" gate "naive" and facilitated receptor states cannot be resolved electrophysiologically, while the associated domain movements can still be detected. In principle, the mechanism of the hinge (gate movement measured by fluorescence) does not change when the door (gate, measured electrophysiologically) has a hole.

5) Do the F change of R-Geco (Figure 6B) and FRET change by mNeonGreen (Figure 6C) disappear in the absence of extracellular Ca²⁺?

R-GECO is a widely used Ca²⁺-sensor and fluorescence changes were not expected in the absence of extracellular Ca²⁺. Since the FRET approach was discontinued, these control experiments were not performed.

Reviewer #3 (Recommendations for the authors):

1. Please cite this paper when talking about eRF-E55D as well, https://doi.org/10.7554/eLife.37248.

Thank you, done (line 103).

2. The 3'-terminal CCA-sequence is already in the pANAP plasmid. More elaboration on how it augmented the translation is needed to make it clear to readers.

The 3’-terminal CCA sequence is important for aminoacylation and translation. The pANAP sequence described in (Chatterjee and Guo, 2013) in the Supporting Information S2 encoding the tRNA lacks a 3’-CCA. This sequence might be dispensable, as it would likely be added by X. laevis oocyte-endogenous tRNA-nucleotidyltransferases. However, we decided to add it, to provide a functional tRNA ready for aminoacylation by the ANAP-specific tRNA synthetase already upon cytoplasmic injection. To avoid misunderstanding, we replaced "augmented" by "provided".

3. Additional FRET experiments might be out of the scope of this paper, but please add more discussion and references regarding this possibility in the paper.

We included several references to illustrate the potential of ANAP as a FRET partner more clearly (lines 328ff).

4. One area that could be improved on is the uncoupled "ballast" domain. How might GTP/GDP and dinuclear Zn2+-binding rearrange this domain? Can pharmacological manipulations be used to change intracellular GTP/GDP or Zn2+ levels to see how Anap labeled in this domain changes its fluorescence differently? Could the palmitoylation of the anchor domain play a role for uncoupling the "ballast' domain and for the current facilitation?

We are not aware of any protocols to change intracellular GTP or Zn²⁺ levels in a time scale that is amenable to our VCF analysis. As described above, we analyzed the Cys-free mutants that are likely not palmitoylated. These analyses confirmed the importance of palmitoylation for the facilitation process and are now included (see Essential revision point 1). However, expression of these mutants was too low for VCF analysis and no obvious effects of palmitoylation on fluorescence amplitude or kinetics could be detected. This is now mentioned in lines 190-205.

5. Line 634, should be "artifacts"?

Thank you, corrected!

https://doi.org/10.7554/eLife.82479.sa2

Article and author information

Author details

Anna Durner

Walther Straub Institute of Pharmacology and Toxicology, Faculty of Medicine, LMU Munich, Munich, Germany

Contribution
Conceptualization, Formal analysis, Investigation, Methodology, Writing - original draft, Writing - review and editing

Competing interests
No competing interests declared

"This ORCID iD identifies the author of this article:" 0000-0002-0993-8869
Ellis Durner

Lehrstuhl für Angewandte Physik and Center for Nanoscience, LMU Munich, Munich, Germany

Contribution
Conceptualization, Resources, Formal analysis, Methodology, Writing - original draft, Writing - review and editing

Competing interests
No competing interests declared

"This ORCID iD identifies the author of this article:" 0000-0002-4461-9257
Annette Nicke

Walther Straub Institute of Pharmacology and Toxicology, Faculty of Medicine, LMU Munich, Munich, Germany

Contribution
Conceptualization, Supervision, Funding acquisition, Writing - original draft, Project administration, Writing - review and editing

For correspondence
annette.nicke@lrz.uni-muenchen.de

Competing interests
No competing interests declared

"This ORCID iD identifies the author of this article:" 0000-0001-6798-505X

Funding

Deutsche Forschungsgemeinschaft (335447717 - SFB 1328, Project A15)

Annette Nicke

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

This work was supported by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation, Project-ID: 335447717 - SFB 1328, A15). We thank Luis Pardo, Kerstin Dümke, and Monika Haberland for providing Xenopus laevis oocytes.

Senior Editor

Richard W Aldrich, The University of Texas at Austin, United States

Reviewing Editor

Stephan A Pless, University of Copenhagen, Denmark

Reviewer

Gucan Dai, Saint Louis University, United States

Publication history

Received: August 5, 2022
Preprint posted: September 1, 2022 (view preprint)
Accepted: January 3, 2023
Accepted Manuscript published: January 4, 2023 (version 1)
Version of Record published: January 20, 2023 (version 2)

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This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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Anna Durner
Ellis Durner
Annette Nicke

(2023)

Improved ANAP incorporation and VCF analysis reveal details of P2X7 current facilitation and a limited conformational interplay between ATP binding and the intracellular ballast domain

eLife 12:e82479.

https://doi.org/10.7554/eLife.82479