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2 The abbreviations used are:
SERCAsarco-endoplasmic reticulum calcium ATPase2-color SERCASERCA with YFP fused to the nucleotide-binding domain and Cer fused to the actuator domainAAAtriple D426A/E429A/E435A mutant of SERCARRruthenium redCercerulean fluorescent proteinE1SERCA enzyme conformation with high-affinity Ca2+ transport sites oriented toward the cytoplasmE2SERCA enzyme conformation with low-affinity Ca2+ transport sites oriented toward the lumenERendoplasmic reticulumIonoionomycinMDmolecular dynamicsPCAprincipal component analysisRyRryanodine receptorTGthapsigarginX-RhodX-Rhod-1/AMYFPenhanced yellow fluorescent proteinTMtransmembranePDBProtein Data BankAMP-PCP5′-adenylyl-β,γ-imidodiphosphate.
The conformational changes of a calcium transport ATPase were investigated with molecular dynamics (MD) simulations as well as fluorescence resonance energy transfer (FRET) measurements to determine the significance of a discrete structural element for regulation of the conformational dynamics of the transport cycle. Previous MD simulations indicated that a loop in the cytosolic domain of the SERCA calcium transporter facilitates an open-to-closed structural transition. To investigate the significance of this structural element, we performed additional MD simulations and new biophysical measurements of SERCA structure and function. Rationally designed in silico mutations of three acidic residues of the loop decreased SERCA domain–domain contacts and increased domain–domain separation distances. Principal component analysis of MD simulations suggested decreased sampling of compact conformations upon N-loop mutagenesis. Deficits in headpiece structural dynamics were also detected by measuring intramolecular FRET of a Cer–YFP–SERCA construct (2-color SERCA). Compared with WT, the mutated 2-color SERCA shows a partial FRET response to calcium, whereas retaining full responsiveness to the inhibitor thapsigargin. Functional measurements showed that the mutated transporter still hydrolyzes ATP and transports calcium, but that maximal enzyme activity is reduced while maintaining similar calcium affinity. In live cells, calcium elevations resulted in concomitant FRET changes as the population of WT 2-color SERCA molecules redistributed among intermediates of the transport cycle. Our results provide novel insights on how the population of SERCA pumps responds to dynamic changes in intracellular calcium.
SERCA with YFP fused to the nucleotide-binding domain and Cer fused to the actuator domain
AAA
triple D426A/E429A/E435A mutant of SERCA
RR
ruthenium red
Cer
cerulean fluorescent protein
E1
SERCA enzyme conformation with high-affinity Ca2+ transport sites oriented toward the cytoplasm
E2
SERCA enzyme conformation with low-affinity Ca2+ transport sites oriented toward the lumen
ER
endoplasmic reticulum
Iono
ionomycin
MD
molecular dynamics
PCA
principal component analysis
RyR
ryanodine receptor
TG
thapsigargin
X-Rhod
X-Rhod-1/AM
YFP
enhanced yellow fluorescent protein
TM
transmembrane
PDB
Protein Data Bank
AMP-PCP
5′-adenylyl-β,γ-imidodiphosphate.
is the ion transporter responsible for sequestering calcium in the sarcoplasmic reticulum (SR) and endoplasmic reticulum (ER). Mutations in the skeletal muscle isoform SERCA1a cause Brody myopathy, with impaired muscle relaxation (
). Alterations in cardiac SERCA2a expression, activity, and regulation have been linked to cardiovascular diseases such as heart failure, hypertrophy, and senescence (
Differential changes in cardiac phospholamban and sarcoplasmic reticular Ca2+-ATPase protein levels: effects on Ca2+ transport and mechanics in compensated pressure-overload hypertrophy and congestive heart failure.
Calcium upregulation by percutaneous administration of gene therapy in patients with cardiac disease (CUPID 2): a randomised, multinational, double-blind, placebo-controlled, phase 2b trial.
). Thus, there is a critical unmet need in the development of small-molecule therapies based on modulation of endogenous SERCA function.
SERCA is composed of four major domains. The cytosolic headpiece consists of an actuator (A) domain, a nucleotide-binding (N) domain, and an autophosphorylation (P) domain. The spatial arrangement of these domains changes during phosphoryl transferase steps in the catalytic cycle, thereby altering the orientation and affinity of the Ca2+-transport sites in the transmembrane (TM) domain (
). SERCA transports two Ca2+ ions per ATP molecule hydrolyzed, with the formation of a phosphoenzyme intermediate (phospho-Asp), and thus is classified as a P-type ATPase (
). The catalytic cycle of P-type pumps was first identified for the Na+/K+-ATPase and is referred to as the Post-Albers transport mechanism, where the Ca2+-transport sites show alternating access (cytosolic and luminal) and alternating affinity (high and low, respectively).
To investigate the rearrangement of SERCA cytosolic domains and identify key structural determinants we previously performed a computational study of SERCA headpiece motions (
). The analysis indicated that a specific structural feature of the N-domain, the Nβ5–β6 loop, facilitates the transition of the SERCA headpiece from an open arrangement of domains to a more compact architecture. This short β-loop is composed of residues 426–436 (426DYNEAKGVYEK436) and contains three negatively-charged amino acids (Asp-426, Glu-429, and Glu-435) that form salt bridges and hydrogen bonds with basic and polar residues on the surface of the A-domain. These interactions help initiate domain–domain contact and eventually induce SERCA headpiece closure. When these three negative charges were mutated in silico to Ala, contacts between the A- and N-domains were decreased, and headpiece dynamics were altered in short MD trajectories. This previous finding suggested that the three negatively-charged residues of the Nβ5–β6 loop are important for SERCA headpiece closure, and we predicted that these residues are key determinants of transport function.
In the present study, we tested this hypothesis with longer MD simulations and additional trajectory analyses, plus we performed physical experiments including functional assays of N-loop mutants. We also measured FRET as an index of overall headpiece conformation. These experiments exploited a “2-color SERCA” construct consisting of fluorescent protein tags fused to the N- and A-domains of cardiac SERCA2a (
). We previously used this 2-color construct as an intramolecular biosensor in high-throughput screening assays to identify small-molecule effectors that affect pump structure and activity (
). Here the 2-color construct was used to report the redistribution of the population of SERCA pumps among different conformational states and to directly test the proposed role of the Nβ5–β6 loop in the structural transitions of the Ca2+ transport cycle.
Results
All-atom molecular dynamics simulations of SERCA mutants
Our previous MD simulations of SERCA structural dynamics predicted that the open-to-closed transition of the SERCA cytosolic headpiece is facilitated by electrostatic and hydrogen bond interactions between basic and polar residues in the A-domain and acidic residues in the N-domain Nβ5–β6 loop (
). The contacts help to pierce a structured and poorly diffusible water layer between the domains. These residue–residue interactions also support a stable, ordered arrangement of the N- and A-domains during the transition from an open to a closed headpiece conformation. In silico triple mutation of the three negatively-charged loop residues to Ala abolished these stabilizing interactions and decreased the likelihood of spontaneous transition of the SERCA headpiece from an open architecture to a closed, compact structure, as assessed by 40-ns MD simulations (
). Here we extended these simulations to 100 ns, ran new simulations of single-point mutants of the SERCA Nβ5–β6 loop, and performed additional structural analyses on all trajectories. Fig. 1A shows the starting X-ray crystal structure of SERCA used for all-atom MD simulations (PDB accession code 1SU4) (
), including the A-, N-, P-, and TM-domains, plus the Nβ5–β6 loop of the N-domain and its three acidic residues, highlighted in orange. For triple mutant AAA (D426A/E429A/E435A) or single substitutions of Nβ5–β6 loop residues, we observed similar root mean square deviation trajectories in N-domain motions and insignificant differences in residue-specific root mean square fluctuations, and the angular autocorrelation of the cytoplasmic domains was not detectably different (data not shown); thus, the overall structure and dynamics of SERCA are not disrupted by single or triple mutations. Covariance matrices were only modestly different for WT- and AAA-SERCA, although there was a decrease in negatively correlated motions of the N- and A-domains (Fig. 1B, dotted box outlines) for AAA. The most pronounced effect of Nβ5–β6 loop mutation observed in the present MD simulations is a reduction in contacts between the N- and A-domains (Fig. 1, C and D). In particular, substitution of three Nβ5–β6 loop residues (Asp-426, Glu-429, and Glu-435) to Ala decreased domain–domain contacts (Fig. 1, C and D, AAA), resulting in greater separation of the domains over the course of the trajectories (Fig. 1E, AAA). Fig. 1F reveals the general trend of a negative dependence of domain separation distance on the number of domain–domain contacts, whereby WT shows a short distance between N- and A-domains stabilized by a large number of contacts, whereas AAA shows a large domain–domain separation and fewer domain–domain contacts. Single-point mutants yielded intermediate values or were comparable with WT (Fig. 1, C–F). The data are consistent with the proposed role of the three acidic N-loop residues in the open-to-closed structural transition (
Figure 1MD simulations of SERCA structural dynamics. For C–E, WT–SERCA (black), AAA (red), D426A (blue), E429A (green), E435A (pink) are shown. Data represent average of 6 MD run productions. A, SERCA starting X-ray structure of PDB 1SU4 for simulations showing the actuator (A), nucleotide-binding (N), phosphorylation (P), and TM domains. The Nβ5–β6 loop is highlighted in orange, and three negatively charged residues Asp-426, Glu-429, and Glu-435 are labeled in the magnified inset. B, covariance matrices Cα atoms for WT–SERCA (upper left) and AAA–SERCA (lower right). Covariance analysis of WT–SERCA residue dynamics as measured from Cα revealed positively (red) and negatively (blue) correlated motions. Dotted boxes highlight regions of covariance of the N- and A-domains. For AAA–SERCA, covariance analysis indicated similar global dynamics yet reduced anti-correlated N- and A-domain motions compared with WT–SERCA. Data show representative of 6 MD runs. C, number of contacts between the N- and A-domains during MD trajectories. D, quantification of results in C. E, AAA–SERCA shows an increase in N- to A-domains separation distance compared with WT-SERCA. F, negative correlation of separation distance on domain–domain contacts, from results in D and E. G, first and second principal components of SERCA domain motions. H, relative sampling of the top two principal components by WT–SERCA (black) and AAA–SERCA (red) trajectories. Each point represents a conformation extracted from the MD trajectories at an interval of 0.1 ns. For comparison, gray dots represent X-ray structures with open (1SU4), closed (1VFP), and intermediate (3W5B) headpiece conformations.
Comparison between structural ensembles of WT–SERCA and AAA–SERCA
We performed principal component analysis (PCA) of the ensembles of WT– and AAA–SERCA trajectories (n = 6 each) and analyzed how the ensemble trajectory of each construct sampled the first two principal components. The most dominant motion PC1 (48% of all motions) was SERCA headpiece opening/closing, and the second major component PC2 (15% of all motions) was twisting of the cytosolic domains. Fig. 1G summarizes the positive and negative deflections of the A- and N-domains along components 1 and 2, with the origin (0, 0) representing the starting equilibrated structure. With respect to PC1, WT-SERCA sampled values ranging from a minimum of −36 Å (closed) to a maximum of +25 Å (open) (Fig. 1H, black). AAA–SERCA trajectories (Fig. 1H, red) showed a similar range along PC1 (minimum of −30 Å and maximum of +31 Å), but with relatively more population of open structures at the expense of closed conformations. This result is consistent with the observed increased minimum N- to A-domain–domain distance for AAA compared with WT (Fig. 1, E and F). WT showed several trajectories that populated the extremes of the PC2 axis (twisting of the N- and A-domains) (Fig. 1H), ranging from −27 to +23 Å, which is 56% greater than the range of AAA–SERCA along PC2 (from −14 to +18 Å). Thus, the AAA mutant showed smaller twisting motions of the N- and A-domains (Fig. 1G). Overall, PCA indicates that the AAA loop mutations decrease the range of motions of SERCA cytosolic domains (with respect to PC2), and shift the population of structures toward a more open architecture (with respect to PC1).
The role of the Nβ5–β6 loop in SERCA ATPase and Ca2+-transport activities
To determine the functional significance of the Nβ5–β6 loop in Ca2+-activated ATP hydrolysis, we prepared ER microsomes from HEK-293 cells expressing 2-color WT–SERCA or AAA–SERCA (Fig. 2, A–C). Fig. 2A shows an example experiment measuring Ca2+-activated ATP hydrolysis by SERCA constructs. The apparent Ca2+ affinity (KCa) of AAA–SERCA was not significantly different from that of WT–SERCA, yielding KCa values of 7.0 ± 0.2 and 7.1 ± 0.3, respectively (n = 4, p = 0.61) (Fig. 2B). However, the maximal activity of AAA–SERCA was decreased by 63 ± 5% (n = 4, p = 0.015) (Fig. 2C). This result is compatible with the previous finding by the Inesi lab that the single-point mutation of Asp-426 to Ala decreases SERCA ATPase activity similarly by 61% (
). Overall, the data demonstrate that the SERCA cycling rate is decreased by mutation of acidic loop residues, yet the mutated transporter still retains the ability to perform Ca2+-activated ATP hydrolysis with the same Ca2+ affinity as WT–SERCA.
Figure 2N-domain β5–β6 loop triple mutation decreases SERCA function.A, calcium-dependent ATPase activity of cells expressing WT– or AAA–SERCA, or nontransfected cells (Ctrl). B, Ca2+ sensitivity of ATPase activity (KCa), as in A, p = 0.61. C, maximal Ca2+-dependent ATPase rate (Vmax), as in A, *, p = 0.015. D, triple-transfected cells: SERCA, RyR, and R-CEPIA1er. ER Ca2+ was depleted by caffeine (Caf) addition, followed by ER Ca2+ store recovery in the presence of RyR blocker ruthenium red (RR). E, maximal Ca2+ uptake rate, as in D. *, p = 0.016 for AAA versus WT and p = 0.002 for Ctrl versus WT. F, maximal ER Ca2+ load, as in D. *, p = 4 × 10−5versus WT.
To determine the rate of Ca2+ transport by the AAA mutant, we quantified intracellular Ca2+ uptake in the ER of HEK-293 cells permeabilized with saponin (Fig. 2, D–F). For each cell, Cer-SERCA expression (WT or AAA) was determined from the emission intensity of Cer fusion tag. To provide pharmacological control over ER Ca2+ content, cells were co-transfected with the cardiac ryanodine receptor (RyR), an SR Ca2+ channel that opens in response to application of caffeine (
). The combination of a high level of Ca2+-transport activity from exogenous Cer-SERCA together with the ER Ca2+-load dependence of the exogenous RyR opening (
) results in spontaneous Ca2+ release events, followed by refilling of the ER Ca2+ stores by SERCA (Fig. 2D: 8 release/uptake events between t = 0–15 s). Cells expressing AAA–SERCA showed similar average basal ER Ca2+ content compared with WT, with few spontaneous Ca2+ release events (Fig. 2D). Spontaneous Ca2+ release was not observed in nontransfected cells (Ctrl) (Fig. 2D), as these cells express a low amount of native SERCA and lack endogenous RyR (
To determine the Ca2+ uptake rate by SERCA pumps, caffeine (10 mm) was applied to empty the ER Ca2+ stores (Fig. 2D, Caf). Once caffeine was washed out, the RyR inhibitor ruthenium red (10 μm) was applied to stop Ca2+ release and allow measurement of the rate of luminal [Ca2+]ER recovery. At the end of each experiment, the R-CEPIA1er signal was calibrated by addition of the Ca2+ ionophore ionomycin (Iono) (
). ER Ca2+ recovery, monitored by [Ca2+]ER accumulation, was analyzed to determine the maximum ER Ca2+ uptake rate (i.e. SERCA transport rate) and maximum ER Ca2+ load. ER Ca2+ uptake in cells expressing WT–SERCA was determined to be 0.14 ± 0.02 mm/s, a 47-fold increase over control cells (0.003 ± 0.0005 mm/s) (Fig. 2E). AAA–SERCA Ca2+ uptake rate was 0.05 ± 0.007 mm/s, a 3-fold reduction in Ca2+ transport rate versus WT. Furthermore, AAA–SERCA generated a ∼25% lower maximal ER Ca2+ load. The differences in Ca2+ uptake rate and ER maximal load were not due to differential expression of WT– and AAA–SERCA, because the Cer fluorescence emission was similar for these two groups: 73 ± 5.0 arbitrary units of WT and 86 ± 11 arbitrary units in AAA–SERCA. The localization of 2-color SERCA was not significantly altered by the AAA mutation, as determined from confocal microscopy. Moreover, we observed similar levels of SERCA protein in ER microsomal preparations as evaluated by comparing exogenous Cer-labeled SERCA with endogenous SERCA by Western blotting. In HEK-293 cell microsomes, exogenous WT–SERCA expression was 68 ± 7% of total SERCA, whereas AAA-SERCA expression was 78 ± 7% of total SERCA. Ponceau staining of the blot also indicated similar amounts of WT– and AAA–SERCA, 6.8 ± 0.6 and 6.5% ± 0.7% of total protein, respectively. The data suggest that translation and localization of SERCA in the ER membrane was similar for WT and AAA. We conclude that AAA–SERCA exhibits lower Ca2+ transport activity (65% less) than WT–SERCA (Fig. 2, D and E), which is consistent with the decreased ATPase rate by AAA–SERCA (63% inhibition) relative to WT (Fig. 2, A and C).
Quantification of 2-color SERCA FRET in ER microsomes from HEK-293 cells
We have previously used intermolecular FRET to quantify SERCA regulatory interactions with phospholamban and sarcolipin (
Phospholamban oligomerization, quaternary structure, and sarco(endo) plasmic reticulum calcium ATPase binding measured by fluorescence resonance energy transfer in living cells.
). The latter experiments utilized a doubly labeled SERCA with fluorescent proteins fused to the N- and A-domains (2-color SERCA). Here we prepared microsomal membranes from cells expressing WT 2-color SERCA to quantify the FRET response to Ca2+ binding using confocal fluorescence microscopy.
Fig. 3A shows that FRET increased with Ca2+ concentration, with an EC50 of 1.25 ± 0.22 μm and a Hill coefficient (n) of 0.76. The apparent lack of cooperativity is compatible with previous studies that suggest the E2–E1 structural transition of the cytosolic headpiece is complete after binding of the first Ca2+ to transport site I (
Effects of phospholamban transmembrane mutants on the calcium affinity, maximal activity, and cooperativity of the sarcoplasmic reticulum calcium pump.
). Overall, the data are consistent with our previous observation that ionophore treatment of HEK-293 cells expressing 2-color SERCA caused accumulation of 2-color SERCA in a high FRET state over the course of a few minutes (
Figure 3Ligand-induced control of SERCA structure in ER microsomes.A, WT 2-color SERCA shows increased FRET with increasing Ca2+, as detected by confocal fluorescence microscopy. Black error bars represent S.E., whereas gray bars represent S.D. (n = 6 experiments). B, FRET of WT 2-color SERCA stabilized in key enzymatic states. *, p ≤ 0.008 compared with H+. C, calculated FRET distances compared with distances between the fluorescent protein fusion sites measured from X-ray crystal structures. Select structures are labeled for comparison, other data are identified in Table 1.
). Importantly, whereas the state designations applied here are widely used in the field to describe the biochemical states stabilized by particular conditions, it is likely that significant structural heterogeneity exists for all ligand-stabilized biochemical states (
). For example, our previous time-resolved fluorescence measurements have shown that SERCA bound to the inhibitor TG can sample at least two major conformations (
). The present measurements capture the average FRET of the population ensemble.
We observed generally low FRET for 2-color SERCA in HEK-293 microsomes in solution conditions under which E2 conformers (low Ca2+ affinity) are expected to predominate (Fig. 3B). For example, E2 (protonated) SERCA yielded ∼10% FRET, as did E2–thapsigargin (E2–TG), a potent inhibitor that locks SERCA in the calcium-free E2 state (
). E2–AlF4− and E2–Vi biochemical intermediates (analogs of the E2P phosphoenzyme intermediate) also showed low FRET (∼8%) (Fig. 3B). In contrast, we observed generally high FRET (∼15%) for 2-color SERCA in microsomes in solution conditions under which E1 conformers (high Ca2+ affinity) are expected to predominate, such E1–ATP, E1–2Ca, 2Ca–AMP-PCP, and E1–2Ca–ADP–AlF4− (Fig. 3B). The FRET values for each state are summarized in Table 1, and these results provide the basis for quantitative analysis of the population distribution of SERCA in HEK-293 cells (see next section). Fluorescent protein separation distances (calculated from measured FRET efficiency) correlated well with distances measured from X-ray crystal structures (Fig. 3C), with the exception of the E1–2Ca crystal structure (PDB code 1SU4). We conclude that SERCA E2 conformers have more open headpiece structures (lower FRET), whereas E1 conformers have more closed headpiece structures (higher FRET).
For initial investigation of SERCA structural changes in response to Ca2+ elevations in live cells, we quantified intramolecular FRET of 2-color SERCA by widefield fluorescence microscopy. Compared with WT, 2-color AAA–SERCA showed decreased basal FRET (Fig. 4A) in live HEK-293 cells, indicating a more open headpiece architecture compared with WT. We regard this result as consistent with our MD simulations, which showed fewer contacts between N- and A-domains for AAA-SERCA (Fig. 1D). To a lesser degree, the single-point mutations of acidic Nβ5–β6 loop residues also reduced FRET in live HEK-293 cells (Fig. 4A). The more moderate phenotypes of D426A, E429A, and E435A are also compatible with MD simulations, which showed domain contacts of the point mutants are intermediate between WT and AAA, or close to WT (Fig. 1D).
Figure 4Ca2+-dependent redistribution of SERCA conformers in live cells.A, FRET of 2-color SERCA constructs expressed in basal HEK-293 cells, as detected by acceptor-sensitized fluorescence microscopy, *, p ≤ 0.005. B, TG-induced E2 conformers of WT 2-color SERCA and the four mutant constructs show reduced FRET in HEK-293 cells, whereas DMSO vehicle-only control (Ctrl, dark blue) had no significant effect. C, addition of the Ca2+ ionophore Iono resulted in a biphasic FRET response (1 = quick decrease, 2 = slow recovery) for WT and single-point mutants, as detected using epifluorescence microscopy. AAA–SERCA (red) showed only the first phase: a quick decrease in FRET. Data represent the average of n = 6–25 cells for each condition.
), and here we found that all of the mutants, including AAA–SERCA, responded normally to this SERCA inhibitor (Fig. 4B). Increasing intracellular Ca2+ by application of ionomycin to cells expressing WT 2-color SERCA resulted in a biphasic response (Fig. 4C, black trace), with a rapid decrease in FRET (phase 1), followed by a slower increase in FRET (phase 2), which recovers to around the initial high FRET value (pre-Iono). Interestingly, the point mutants all showed a normal two-phase FRET response to Ca2+ (like WT) (Fig. 4C, blue, green, and magenta traces). However, mutation of all three acidic loop residues (AAA-SERCA) abolished the slow FRET increase in phase 2 (Fig. 4C, red trace). Proposed mechanistic origins of the two-phase response are detailed under “Discussion.”
Overall, the widefield fluorescence microscopy data are consistent with the MD simulation analyses, showing that mutation of one negative residue is moderately tolerated, but that mutation of three acidic residues from the Nβ5–β6 loop impairs N- to A-domain contacts (Fig. 1D), thereby decreasing the likelihood of forming a more compact headpiece conformation. Importantly, the observation that AAA–SERCA can still respond to TG in live cells (Fig. 3B) indicates that the protein structure is intact and TG-induced changes in headpiece structural dynamics are preserved after loop mutation. Thus, we propose that the impaired second-phase of Ca2+ response and the reduced activity of AAA–SERCA are not due to a gross structural defect (such as misfolding), but instead due to the lack of the N-domain structural determinant that is responsible for the stable interaction with the A-domain during the SERCA headpiece transition from an open-to-closed conformation. These FRET experiments serve as the foundation for assessing the redistribution of SERCA calcium pump conformers during dynamic intracellular Ca2+ signaling.
SERCA structural dynamics in response to changes in intracellular Ca2+ concentration
We have previously quantified FRET in cardiac myocytes (
), but motion artifacts of actively contracting cells make quantification of dynamic changes in fluorescence challenging. As an alternative, we reconstituted aspects of muscle cell Ca2+ handling with co-expression of RyR2 and SERCA2a in live HEK-293 cells, and subjected the proteins to confocal microscopy.
In these experiments, Ca2+ in the cytosol or ER lumen was monitored with X-Rhod (X-Rhod-1/AM) or R-CEPIA1er, respectively. SERCA conformational changes were quantified by excitation of Cer at 458 nm and observed as anti-correlated changes in the fluorescence intensities of Cer and enhanced yellow fluorescent protein (YFP) (Fig. 5A). The ratio of YFP/Cer fluorescence was taken as a measure of relative intramolecular FRET in the SERCA headpiece (Fig. 5, B, C, and E–L). Based on microsomal membrane experiments that showed high FRET for E1 states (Fig. 3B), and the eventual accumulation of a high FRET state after ionomycin treatment of cells (Fig. 4C), we anticipated that spontaneous Ca2+ release events would be accompanied by increases in intramolecular FRET. Instead, Ca2+ elevations in the cytosol corresponded to reductions in FRET, as indicated by decreases in the ratio of YFP/Cer fluorescence (Fig. 5B). Likewise, depletions of ER Ca2+ occurred simultaneously with decreases in SERCA intramolecular FRET (Fig. 5C). We noted that the recovery of basal SERCA FRET was complete before full restoration of ER Ca2+ stores, as shown by SERCA returning to the low FRET state in the middle of the sawtooth profile in ER Ca2+ content detected by R-CEPIA1er (Fig. 5C). In contrast, cytosolic Ca2+ elevations were square steps that closely mirrored SERCA FRET depressions (Fig. 5B). The result suggests that the ER continues to fill even after cytosolic Ca2+ is already back to baseline and the population of SERCA has returned to the basal high FRET conformation. This may be due to store-operated Ca2+ entry mechanisms that largely bypass the bulk cytoplasm.
Figure 5SERCA structural dynamics measured by FRET in HEK-293 live cells.A, anti-correlated changes in Cer and YFP fluorescence intensity indicate rhythmic FRET fluctuations in intact cells. B, the ratio of YFP/Cer, as in A, was used as an index of FRET (gray). FRET (bottom panel, gray) was inversely correlated to cytosolic Ca2+, as measured by X-Rhod fluorescence (black, top panel). WT–SERCA FRET decreased during cytosolic Ca2+ elevations due to spontaneous ER Ca2+ release through RyR. C, a decrease in 2-color SERCA intramolecular FRET (bottom trace) occurs simultaneously with depletion of ER Ca2+ stores (top trace). D, quantification of intermolecular FRET between Cer–SERCA and YFP–SERCA using progressive acceptor photobleaching (started at black arrow). Data presented are the mean F/F0 of Cer and YFP fluorescence measured in 6 cells. E, SERCA–SERCA intermolecular FRET did not change in response to ionomycin addition. F, SERCA–SERCA intermolecular FRET did not change with spontaneous Ca2+ release events (top trace). The data indicate that changes in 2-color SERCA FRET are due to changes in intramolecular FRET rather than changes in intermolecular FRET. G, after addition of ionomycin, both cytosolic Ca2+ and WT–SERCA FRET increased, as detected by confocal fluorescence microscopy. H, addition of caffeine (Caf) transiently increased cytosolic Ca2+ and decreased WT–SERCA FRET. I, AAA–SERCA FRET decreased during spontaneous cytosolic Ca2+ elevations. J, in contrast to WT–SERCA FRET, AAA–SERCA FRET decreased with addition of ionomycin. K, AAA–SERCA FRET increased in response to caffeine, similar to WT–SERCA FRET (H). L, addition of ionomycin causes an increase in ER Ca2+ content, but this increase occurs more slowly than the second phase of the observed FRET response of 2-color SERCA. We conclude that the phase 2 FRET increase of 2-color SERCA is not due to saturation of SERCA luminal Ca2+-binding sites (i.e. low affinity E2 orientation).
). Although we detected intermolecular FRET between Cer–SERCA and YFP–SERCA under these experimental conditions (Fig. 5D), intermolecular FRET did not change in response to ionomycin (Fig. 5E) or spontaneous Ca2+ release events (Fig. 5F).
Despite the surprising response to intracellular Ca2+ release, WT–SERCA intramolecular FRET still showed a biphasic response (decrease, then increase) in these confocal microscopy experiments after addition of ionomycin. Fig. 5G compares these contrasting results in a single trace: transient elevations of Ca2+ resulted in transiently decreased FRET, but a sustained increase in Ca2+ after ionomycin addition caused a lagging increase in FRET. As an alternative, we activated the co-expressed RyR with caffeine, and observed transiently increased cytosolic Ca2+ followed by rapid equilibration to a low concentration of cytosolic Ca2+ (Fig. 5H). Remarkably, this event was mirrored by a FRET change (decrease, then increase) that had similar kinetics.
In light of the poor responsiveness of AAA–SERCA to ionomycin-induced Ca2+ influx (Fig. 4C, red), we were also surprised to find AAA–SERCA responded to spontaneous cytosolic Ca2+ elevations (Fig. 5I) with no apparent deficit compared with WT (Fig. 5B), but again, Ca2+ influx after ionomycin addition yielded a sustained decrease in FRET (Fig. 5J) instead of the biphasic FRET response seen for WT (Fig. 5G). The FRET response of AAA–SERCA to caffeine (Fig. 5K) was similar to that of WT–SERCA (Fig. 3H); a transient decrease followed by a sustained increase in FRET.
Discussion
In the present study we aimed to quantify SERCA dynamics during calcium signaling in live cells to examine the role of a discrete structural element that we previously hypothesized was an important determinant of SERCA headpiece structural transitions. Our new experiments confirm and extend our previous results (
). In particular, new MD experiments further defined the structural role of the acidic residues in the Nβ5–β6 loop, which is to establish inter-domain contacts that determine the range of domain movements and facilitate headpiece closure. The functional significance of this structural element is indicated by in vitro ATPase assays and live cell Ca2+ uptake assays, which showed that mutations of key loop residues reduced ATPase activity and Ca2+ transport kinetics. In addition, simultaneous FRET and Ca2+ measurements provided insight into the dynamic redistribution of SERCA conformers in the physiologically relevant context of the cell (Fig. 5). To interpret the results of Ca2+ dynamics experiments, we also quantified the average headpiece conformation of ligand-stabilized enzymatic intermediates (Fig. 3B), which was useful for gauging the average conformation of the population of SERCA molecules for each biochemically defined intermediate state.
Fig. 6A shows a simplified Post-Albers reaction cycle, with high Ca2+-affinity states highlighted in yellow and low Ca2+-affinity states highlighted in green. The relative distribution of SERCA molecules among the conformational states depends on whether ligands such as Ca2+ are abundant or limiting. Importantly, when ligands are not limiting, the distribution of states depends on the relative kinetics of partial reactions (
), with an increased population of states preceding slow steps. Major physiological states and transitions are shown in black (Fig. 6A), plus an additional nonphysiological state (TG-inhibited) and an alternate pathway (Ca2+ binding prior to ATP) are shown in gray. States with similar headpiece conformations are grouped in blue boxes annotated with the FRET efficiency observed for those conformations in steady-state experiments (Fig. 3). In this context, we may interpret the observed changes in FRET in live cells.
Figure 6Redistribution of SERCA conformers during Ca2+ signaling.A, a simplified Post-Albers cycle. Blue boxes enclose states with similar intramolecular FRET efficiency. B, schematic diagram of changes in cytosolic Ca2+ (red) and changes in FRET (gray) as the population of SERCA redistributes among structural states, with the predominant state shown in blue. The WT FRET response to ionomycin shows two phases (phases 1 and 2), whereas AAA FRET shows only a decrease in FRET (dotted line).
Intracellular Ca2+ signaling and SERCA conformational changes
Redistribution of SERCA conformers in response to changes in the concentration of cytosolic Ca2+ during signaling is summarized in the schematic diagram of Fig. 6B. When cells are at rest (e.g. nonstimulated HEK-293 cells or noncontracting myocytes), Ca2+ is low (∼100–200 nm) and ATP is saturating (3–5 mm), so the major populated conformation of SERCA is E1–ATP. Pre-bound ATP shifts the Ca2+ binding E2–E1 (apo) population toward the high affinity E1 (apo)–ATP conformer ready to bind Ca2+ (
). This state exhibited high FRET in vitro, and indeed we observed high FRET for WT 2-color SERCA in HEK-293 cells in the basal state. Then, when Ca2+ is released from intracellular stores into the cytosol, FRET is transiently decreased (Fig. 6B). This FRET change occurs as a consequence of redistribution of the population of transporters to more open headpiece conformations. Specifically, at high Ca2+ the pumps are continuously cycling, and all states of catalytic cycle are populated, with a relative build-up of E1P–2Ca and E2 conformers before the slow interconversion step of the transport sites, plus the E2P conformer before the slow phosphoenzyme hydrolysis and release (Fig. 6A) (
The ATP-induced change of tryptophan fluorescence reflects a conformational change upon formation of ADP-sensitive phosphoenzyme in the sarcoplasmic reticulum Ca2+-ATPase: stopped-flow spectrofluorometry and continuous flow-rapid quenching method.
Assembly of a Tyr-122 hydrophobic cluster in sarcoplasmic reticulum Ca2+-ATPase synchronizes Ca2+ affinity reduction and release with phosphoenzyme isomerization.
). Therefore, when Ca2+ increases to micromolar concentrations the population shifts from being predominantly E1–ATP (high FRET) to a predominant mixture of E1P–2Ca (high FRET) and E2 (low FRET) (Fig. 6A) (
The ATP-induced change of tryptophan fluorescence reflects a conformational change upon formation of ADP-sensitive phosphoenzyme in the sarcoplasmic reticulum Ca2+-ATPase: stopped-flow spectrofluorometry and continuous flow-rapid quenching method.
Assembly of a Tyr-122 hydrophobic cluster in sarcoplasmic reticulum Ca2+-ATPase synchronizes Ca2+ affinity reduction and release with phosphoenzyme isomerization.
). The result of accumulation of the low FRET E2 population decreased overall FRET. Such conditions occur during contractions of cardiac or skeletal muscle cells, or in nonmuscle cells during intracellular Ca2+ waves (as in Figs. 2D and 5B). Termination of Ca2+ release and restoration of basal Ca2+ returns the pumps to the resting condition, in which the majority conform to E1–ATP. Thus, the redistribution of SERCA conformers during Ca2+ signaling is somewhat counterintuitive: the E1–ATP state prevails during periods of low Ca2+, and the E2 state population increases at high Ca2+.
Likewise, bottlenecks in catalytic cycle account for the rapid decrease in FRET immediately after ionomycin addition to HEK-293 cells (Fig. 6B, phase 1) as the population of transporters redistributes to a mixture of E1P/E2. This is the point where the WT and mutant SERCA diverge: Fig. 6B shows that WT–SERCA FRET rebounds (phase 2), whereas AAA–SERCA remains in predominantly low FRET conformations (Fig. 6B, dotted line). The phase 2 increase in FRET for WT-SERCA was remarkable because cytosolic Ca2+ remained elevated at millimolar concentrations for the remainder of the experiment. Several possible mechanisms could account for this result. We considered the possibility of alkalinization of the cell through Ca2+/H+ exchange by ionomycin (
). If this is the origin of the apparent second phase of the FRET change it must be due specifically to a change in SERCA conformation as opposed to a direct effect of pH on the fluorescent protein tags, because control experiments with Cer–SERCA and YFP–SERCA showed no change in intermolecular FRET with ionomycin addition (Fig. 5E). We do not attribute the phase 2 FRET change to saturation of SERCA luminal Ca2+ binding and accumulation of SERCA in the E2–2Ca state, because accumulation of Ca2+ in the ER after ionomycin treatment was much slower than the phase 2 FRET change (Fig. 5L). Another possible mechanism for the phase 2 FRET change is that full activation of SERCA in the ionomycin-treated cells may deplete ATP and increase ADP over the course of several minutes, with consequent accumulation of the population of 2-color SERCA in ATP-free, high FRET conformations. In vitro measurements revealed E1–2Ca and E1–2Ca–ADP–AlF4− to be a high (15%) FRET conformations (Fig. 3B). Finally, SERCA cycling is inhibited at very high (mm) Ca2+ concentrations (
The average conformation at micromolar [Ca2+] of Ca2+-ATPase with bound nucleotide differs from that adopted with the transition state analog ADP: AlFx or with AMPPCP under crystallization conditions at millimolar [Ca2+].
). Thus, we propose that the high FRET observed upon ionomycin-induced Ca2+ release (Figs. 5G and 6A) is due to stabilization of E1–2Ca (branched gray pathway in Fig. 6A). Interestingly, a recent molecular dynamics study of SERCA at supraphysiologic Ca2+ concentration (10 mm) indicated possible Ca2+ binding to the Nβ5–β6 loop (
The failure of AAA–SERCA to undergo a phase 2 redistribution to high FRET states (Fig. 6B, dotted line) illustrates the consequences of mutation of acidic residues in the Nβ5–β6 loop for SERCA dynamics. We observed that the triple mutation AAA impairs SERCA cytosolic headpiece closure in silico and we attribute the lack of a phase 2 FRET change to the decreased kinetics of structural transition from low FRET states (open headpiece) to high FRET states (closed headpiece). Thus, loss of N- to A-domain contacts results in a new rate-limiting step in the SERCA catalytic cycle, the E2 to E1–ATP transition (Fig. 6A). Although the mutant pump is functional (Fig. 2), there is much greater accumulation of the population of cycling transporters in E2 when there is a sustained elevation of Ca2+, so FRET remains low, with no second phase increase.
Importantly, impaired headpiece dynamics did not prevent apparently normal AAA–SERCA FRET changes during spontaneous Ca2+ release events and subsequent Ca2+ re-uptake into the ER lumen (Fig. 5I). In particular, we did not detect any apparent delay in the upstroke of the FRET signal that occurs concomitantly with the decrease in cytosolic Ca2+. This surprising phenomenon may now be understood as another indication that SERCA function is decreased, but not abolished, by loop mutations. Termination of Ca2+ release allows resequestration of Ca2+ by the combined efforts of AAA–SERCA and endogenous SERCA. After withdrawal of Ca2+, flux through the transport cycle ceases, permitting return of AAA–SERCA back to the high FRET basal conformation, E1–ATP. The kinetic delay due to disruption of structural determinants in the loop is not resolvable as a slow FRET transition on this timescale; it is only detectable as a bottleneck step in actively cycling AAA-SERCA.
Comparison with other structural studies
The present results are in harmony with X-ray crystallography of SERCA stabilized by ligands in different biochemical states, with relatively open crystal structures (PDB 1IWO (
), which showed a widely open headpiece conformation. We measured high FRET values for this state in vitro and under conditions that should stabilize this state in cells, consistent with a closed headpiece. Fig. 3C shows the close correlation of the calculated FRET distance and the distance between fluorescent protein fusion sites, as measured from X-ray structures. The regression of these data (excluding outlier 1SU4) yielded a y intercept of 36 Å. We attribute this offset to the additional distance conferred by the difference between the fusion site and the chromaphore at the center of the fluorescent protein β-barrel.
A recent single-molecule FRET study of a related ATPase revealed a similar trend of high FRET between A- and P-domains for E1 conformers and low FRET for A- and P-domains for E2 conformers (
). Although it is difficult to compare FRET measurements taken from different labeling sites, overall the studies are in harmony. Both suggest E1 conformations have a compact, ordered headpiece, whereas E2 states are characterized by an open, disordered architecture.
Summary
The goals of this study were 2-fold. The first was to quantify how the overall conformation of the SERCA cytosolic headpiece changes as the transporter steps through the structural transitions of the catalytic cycle. Second, we sought to test directly the hypothesis that residues in a loop of the SERCA N-domain are key determinants of transport function (
). The present results are compatible with this proposed mechanism, as mutations of the loop residues resulted in altered headpiece dynamics, and functional measurements revealed a consequent decrease in ATP hydrolysis rate and Ca2+ transport. The results support the proposed role of the loop in facilitating SERCA headpiece closure during functional enzymatic cycling. As a discrete structural element, the N-loop may be a worthwhile target for development of small molecules to enhance (
) was used to run WT simulations and to introduce Nβ5–β6 loop mutations. For all constructs, energy minimization was performed using the steepest descent method for 1000 steps. Then the structures were embedded into a POPC lipid bilayer and solvated in a rectangular water box with dimension sizes 130 × 130 × 160 Å. Na+ and Cl− ions were added to the solution to a concentration of 150 mm. The Berendesen method (
) with relaxation time of 0.1 ps was used to increase the temperature of the system to 300 K and reach the pressure of 1 bar. After 1-ns equilibration, the production run was performed in the NPT assemble using the Nose-Hoover thermostat (
) with a relaxation time of 1 ps. Six independent production runs for each WT or mutant construct were started with a different set of assigned velocities at 300 K. The integration time was 2 fs, and atom coordinates were saved every 1 ps. Production runs were carried out for 100 ns (n = 6 for WT– and AAA–SERCA).
) was used for the computational analysis of MD production runs. The N- to A-domain–domain distance was defined as the minimum distance between any atom of the A-domain (residues number 1–40 and 128–241) and any atom of the N-domain (residues 360–603). The number of contacts was calculated with a 4-Å cutoff between atoms of the N- and A-domains. One contact of a N-domain atom with multiple atoms of A-domain was counted as one contact (i.e. instead of multiple contacts). The first 10 ns of MD simulations were considered as equilibration time and not included in structural analyses.
Principal component analysis
To identify the major motions of the SERCA headpiece during MD trajectories, we aligned SERCA structures using the 10-helix TM domain as a reference and used PCA (
)), six MD trajectories of WT–SERCA, and six MD trajectories of AAA–SERCA into a single trajectory. To obtain sets of eigenvectors and eigenvalues corresponding to principal components, we built covariance matrixes of the atomic fluctuations in GROMACS (
). We used a canine SERCA2a construct labeled with Cer on the N terminus and an YFP intrasequence tag inserted before residue 509 in the N-domain for FRET experiments (transient transfection). For ATPase assay experiments, we used an analogous 2-color SERCA construct, with a red fluorescent protein (tagRFP) on the N terminus and an enhanced GFP inserted before residue 509 in the N-domain. The Cer–YFP pair has a Förster distance (R0) of 49.8 Å, and the GFP–tagRFP pair has an R0 of 58.3 Å (
). We introduced Ala mutations in Nβ5–β6 loop residues with QuikChange Lightning Site-directed Mutagenesis Kit (Agilent Technologies, Stratagene, La Jolla, CA) according to the manufacturer's protocol. Single mutations and a triple mutation were made: D426A, E429A, E435A, and D426A/E429/E435A (AAA). Adenoviral vectors of tagRFP–GFP–SERCA (WT and AAA) were produced by the Loyola Cardiovascular Research Institute virus production facility.
HEK-293 cells were cultured in Dulbecco's modified Eagle's medium cell culture medium supplemented with 10% fetal bovine serum (ThermoScientific, Waltham, MA) and transiently transfected using MBS mammalian transfection kit (Agilent Technologies, Stratagene), as described previously (
). The transfected cells were trypsinized (ThermoScientific) and replated onto poly-d-lysine-coated glass bottom chambers and allowed to adhere for 1–2 h prior to imaging.
Wide-field acceptor sensitization fluorescence microscopy to measure SERCA intramolecular FRET
Wide-field fluorescent microscopy was done as described previously (
). Briefly, cells were imaged with an inverted microscope (Nikon Eclipse TE2000-U) equipped with a metal halide lamp and a back-thinned CCD camera (iXon 887: Andor Technology, Belfast, Northern Ireland). For each sample, acquisition of field was performed with a ×60 1.49 N.A. objective with 100 or 150 ms exposure for each channel: Cer, YFP, and FRET. Fluorescence intensity was automatically quantified with a multiwavelength cell scoring application in MetaMorph software (Molecular Devices, Sunnyvale, CA). FRET efficiency was calculated according to E = G/(G + 3.2 × FCer), where G = FFRET − a × FYFP − d × FCer (
), where FFRET, FYFP, and FCer are the matching fluorescence intensities from FRET, YFP, and Cer images, respectively, and G represents FRET intensity corrected for the bleedthrough of the channels. The parameters a and d are bleedthrough constants calculated as a = FFRET/FCer for a control sample transfected with only YFP–SERCA and d = FFRET/FCer for a control sample transfected with only Cer–SERCA. These values were determined to be a = 0.074 and b = 0.70. Apparent probe separation distance (R) was calculated from FRET efficiency (E) according to the relationship (
). The error of the distance measurement was estimated from the standard deviation of repeated FRET measurements.
Ratiometric confocal fluorescence microscopy to measure SERCA intramolecular FRET, HEK-293 cytosolic Ca2+, and HEK-293 ER luminal Ca2+
HEK-293 cells were transiently co-transfected with expression plasmids containing cDNA of GFP-human ryanodine receptor-2 fusion protein (RyR), R-CEPIA1er, and 2-color canine SERCA2a wildtype (WT) or triple loop mutant AAA. Transfected cells were cultured for 24 h and seeded into poly-d-lysine-coated glass-bottom chamber slides in Dulbecco's modified Eagle's medium plus 10% fetal bovine serum. 24 h after seeding, cell culture medium was changed with PBS with Ca2+/Mg2+, and experiments were conducted with a Leica SP5 laser scanning confocal microscope equipped with a ×63 water objective. R-CEPIA1er was excited with the 543-nm line of a He-Ne laser, and emitted fluorescence was measured at wavelength ≥580 nm. 2-color SERCA fluorophores Cer and YFP were excited with the 430 and 514 nm lines of an argon laser, respectively, and emitted fluorescence was measured at wavelengths 485 ± 15 and 537 ± 15 nm, respectively. Images were acquired in line-scan mode for up to 8–12 min with addition of 10 mm caffeine or 100 μm ionomycin at time (t) indicated in the figures. Ionomycin powder (Sigma) was dissolved in DMSO to make a 13.3 mm stock solution, which was used to prepare 2× Iono solution (200 μm) in PBS. The final concentration of DMSO applied to cells was 0.75%. Fluorescence image analysis was performed with ImageJ software (
), cells were incubated with 10 μm X-Rhod (ThermoScientific) for 15 min in PBS (+Ca/+Mg), and then washed twice with PBS (+Ca/+Mg) to remove X-Rhod from the cell culture medium.
HEK-293 cell microsomal membrane preparation
ER microsomal membranes expressing SERCA were isolated from HEK-293 cells infected with adenovirus encoding 2-color WT–SERCA or AAA–SERCA (for ATPase assay) or transfected with 2-color SERCA constructs (FRET measurement) as described (
). Cells were grown to confluence on 150 mm2 dishes for 2 days, washed twice with PBS, harvested by scraping, and pelleted at 1000 × g for 10 min at 4 °C. To prepare cell homogenates, the cell pellets were 1) resuspended in cold homogenizing solution (0.5 mm MgCl2, 10 mm Tris-HCl, pH 7.5, plus EDTA-free complete protease inhibitor mixture (Santa Cruz Biotechnology, Inc., Dallas, TX); 2) disrupted by 10 strokes in a Potter-Elvehjem homogenizer; 3) supplemented with an equal volume of sucrose solution (100 mm MOPS, pH 7.0, 500 mm sucrose, plus EDTA-free complete protease inhibitor mixture); and 4) passed through a 27-gauge needle 10 times. To prepare microsomal membranes, 1) cell homogenates were centrifuged at 1,000 × g for 10 min at 4 °C; 2) the low-speed supernatants were centrifuged at 126,000 × g for 30 min at 4 °C; 3) the high-speed membrane pellets were resuspended in a 1:1 mixture of homogenizing and sucrose solutions; and 4) the membrane suspensions were passed through a 27-gauge needle 10 times. The protein concentration of microsomal membranes were determined with a Pierce BCA assay kit (ThermoScientific). ATPase assay was performed the same day as membrane preparation; otherwise, microsomal membranes were aliquoted, snap-frozen in liquid nitrogen, and stored at −80 °C.
Measuring Ca2+-ATPase activity in ER microsomes from HEK-293 cells
2-Color SERCA ATPase activity was measured in ER microsomes from HEK-293 cells by spectrophotometric determination of the rate of NADH consumption as a function of Ca2+ concentration using the enzyme-coupled activity assay in 96-well plate (
Functional and physical competition between phospholamban and its mutants provides insight into the molecular mechanism of gene therapy for heart failure.
). The time dependence of the absorbance decrease was measured at 340 nm at 25 °C in a PHERAstar FSX microplate reader (BMG Labtech, Cary, NC). Each well contained 3–4 μg of microsomal membranes in 200 μl of assay solution containing 50 mm MOPS, pH 7.0, 100 mm KCl, 5.0 mm MgCl2, 1.0 mm EGTA, 2.5 mm ATP, 0.2 mm NADH, 1 U of pyruvate kinase, 1 U of lactate dehydrogenase, 0.5 mm phosphoenol pyruvate, and 0.7 μg of Ca2+ ionophore (A23187). Chemicals were obtained from Sigma. Free Ca2+ concentrations were calculated using a Ca/Mg/ATP/EGTA calculator from Theo Schoenmakers' Chelator (
where V is the ATPase rate at a specific Ca2+ concentration (pCa), n is the Hill coefficient, pKCa is the apparent Ca2+ dissociation constant, and Vmax was obtained from the fit of the Hill equation at saturating Ca2+ concentrations.
Measuring ER Ca2+ uptake in permeabilized HEK-293 cells
Changes in [Ca2+]ER were measured with laser scanning confocal microscopy (Radiance 2000 MP, Bio-Rad) equipped with a ×40 oil-immersion objective lens (NA = 1.3). R-CEPIA1er was excited with the 543 nm line of a He-Ne laser and fluorescence was measured at wavelengths >600 nm. Fluorescence measurements were acquired in line-scan mode (20 ms per scan; pixel size 0.12 μm). HEK-293 cells co-transfected with GFP-RyR, R-CEPIA1er, and Cer-SERCA were washed in Ca2+-free solution containing 150 mm K-aspartate, 0.25 mm MgCl2, 0.1 mm EGTA, 10 mm HEPES, pH 7.2. The plasma membrane of HEK-293 cells was permeabilized with 0.005% saponin to control cytosolic environment replacement with a saponin-free solution containing 100 mm K-aspartate, 15 mm KCl, 5 mm KH2PO4, 5 mm MgATP, 0.35 mm EGTA, 0.12 CaCl2, 0.75 mm MgCl2, 10 mm phosphocreatine, 2% (w/v) dextran (Mr 40,000), and 10 mm HEPES, pH 7.2 (KOH). Free [Ca2+] and [Mg2+] of this solution were calculated to be 250 nm and 1 mm, respectively. Activation of RyR with caffeine (10 mm) was used to completely deplete [Ca2+]ER. Once caffeine was removed, the RyR inhibitor ruthenium red (RR = 10 μm) was applied to measure the rate of ER Ca2+ uptake. Changes in [Ca2+]ER were calculated by the formula: [Ca2+]ER = Kd × [(F − Fmin)/(Fmax − F)], where F is the R-CEPIA1er fluorescence; Fmax and Fmin are the fluorescence level at 10 mm Ca2+/Iono before and after depletion of ER Ca2+ with caffeine (10 mm), respectively. The R-CEPIA1er Ca2+ dissociation constant (Kd) is 390 μm based on in situ calibrations (
). At the end of each experiment, the R-CEPIA1er signal (Fmax) was calibrated with addition of Iono (100 μm). ER Ca2+ uptake (i.e. SERCA transport rate) was calculated as the time-dependent change of [Ca2+]ER after RyR inhibition on a cell-to-cell basis (mm Ca2+/s), and the maximal ER Ca2+ load was determined for each individual cell (mm Ca2+). The reported Ca2+ uptake rate and maximal ER Ca2+ load were calculated as mean ± S.D.
Intramolecular FRET measurements of 2-color SERCA expressed in ER microsomes from HEK-293 cells
To stabilize SERCA in ligand-stabilized biochemical intermediates, various solutions were prepared by addition of corresponding substrates to the calcium-free base solution, which includes 100 mm KCl, 5 mm MgCl2, 2 mm EGTA, and 10 mm imidazole, pH 7.0. The following ligands were used to prepare specific solutions corresponding to SERCA biochemical state (in parentheses): 100 μm thapsigargin (E2–TG), 3 mm ATP (E1–ATP), 2.1 mm CaCl2 (E1–2Ca) with free [Ca2+]i = 100 μm (
); 2.1 mm CaCl2 and 3 mm nonhydrolyzable ATP analog AMP-PCP (E1–2Ca-AMPPCP); 2.1 mm CaCl2, 3 mm ADP, 3 mm KF, and 50 μm AlCl3 (E1–2Ca–ADP–AlF4−); 0.1 mm orthovanadate (E2–Vi); and 50 μm AlCl3 and 3 mm KF (E2–AlF4−). Chemicals were obtained from Sigma.
To measure SERCA intramolecular FRET in ligand-stabilized biochemical intermediates, 1 μl of membrane preparations (7–10 μg of total protein) was mixed with 9 μl of ligand solution on a coverslip and immediately imaged using confocal fluorescent microscopy, as described above.
Statistical analyses
Data are presented as the mean ± S.D. of n ≥ 3 experiments. All statistical tests were performed using OriginPro 9.1 (OriginLab Corporation, Northampton, MA). Student's t test was used to compare differences between two groups, and one-way analysis of variance was used to compare the difference between three or more groups. One-way analysis of variance was followed by Tukey's post hoc test. A probability (p) value of <0.05 was considered significant. Specific values are provided in figure panels or figure legends.
Author contributions
O. N. R., N. S., and S. L. R. conceptualization; O. N. R. data curation; O. N. R., J. M. A., A. V. Z., and S. L. R. formal analysis; O. N. R., E. B., S. B., and A. V. Z. investigation; O. N. R., N. S., and S. L. R. methodology; O. N. R. and S. L. R. writing-original draft; O. N. R., J. M. A., and S. L. R. writing-review and editing; N. S. and A. V. Z. resources; N. S., A. V. Z., and S. L. R. supervision; S. L. R. funding acquisition; S. L. R. project administration.
Acknowledgments
We are grateful for helpful suggestions from Howard S. Young. This work was also supported by equipment and facilities provided by National Institute of Health “Loyola Research Computing Core” Grant 1G20RR030939. This work used the Extreme Science and Engineering Discovery Environment (XSEDE), which is supported by National Science Foundation Grant ACI-1548562, as well as Stampede and Stampede2 at the Texas Advanced Computing Center (TACC) through XSEDE allocation TG-MCB130108.
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Assembly of a Tyr-122 hydrophobic cluster in sarcoplasmic reticulum Ca2+-ATPase synchronizes Ca2+ affinity reduction and release with phosphoenzyme isomerization.
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This work was supported by the National Institutes of Health Grants HL092321 (to S. L. R) and HL130231 (to A. V. Z.) and the Loyola Cardiovascular Research Institute. The authors declare that they have no conflicts of interest with the contents of this article. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.
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