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Age-dependent diastolic heart failure in an in vivo Drosophila model

  1. Matthew P Klassen  Is a corresponding author
  2. Christian J Peters
  3. Shiwei Zhou
  4. Hannah H Williams
  5. Lily Yeh Jan
  6. Yuh Nung Jan  Is a corresponding author
  1. Howard Hughes Medical Institute, University of California, San Francisco, United States
  2. University of California, San Francisco, United States
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Cite this article as: eLife 2017;6:e20851 doi: 10.7554/eLife.20851

Abstract

While the signals and complexes that coordinate the heartbeat are well established, how the heart maintains its electromechanical rhythm over a lifetime remains an open question with significant implications to human health. Reasoning that this homeostatic challenge confronts all pulsatile organs, we developed a high resolution imaging and analysis toolset for measuring cardiac function in intact, unanesthetized Drosophila melanogaster. We demonstrate that, as in humans, normal aging primarily manifests as defects in relaxation (diastole) while preserving contractile performance. Using this approach, we discovered that a pair of two-pore potassium channel (K2P) subunits, largely dispensable early in life, are necessary for terminating contraction (systole) in aged animals, where their loss culminates in fibrillatory cardiac arrest. As the pumping function of its heart is acutely dispensable for survival, Drosophila represents a uniquely accessible model for understanding the signaling networks maintaining cardiac performance during normal aging.

https://doi.org/10.7554/eLife.20851.001

Introduction

During an average human life span, the heart will undergo 2.8 billion cycles of contraction and relaxation, wherein its four chambers rhythmically beat in a precisely choreographed sequence to efficiently circulate the blood. However, despite our understanding of the molecular basis of the heartbeat (Bers, 2008; Monfredi et al., 2013; Solaro, 2010), societies must confront an increasing failure of this process, which represents an intersection between normal aging and deleterious environmental and genetic factors (Yancy et al., 2013). Previous research has demonstrated that many biophysical properties of the heart are altered over the course of a lifetime (Carrick-Ranson et al., 2012; Cheng et al., 2009; Lakatta et al., 2014). These changes exist structurally, with myocardial fibrosis and arterial stiffening playing a leading role, but also manifest functionally, with alterations in the electrical waveform and calcium handling. However, whether these changes reflect normal senescence or compensatory attempts to maintain function remains an intense area of investigation.

The maintenance of cellular and organismal physiology, termed homeostasis, is essential for all living organisms. To uncover the mechanisms that maintain cardiac function, it has been necessary to develop a systems perspective (Kohl et al., 2010). These efforts have revealed that the heart’s rhythmic behavior is not uniform but exhibits variations that precisely match local mechanical needs (Brutsaert, 1987; Schram et al., 2002). Alterations in the heart’s rhythm also occur during normal aging, where the action potential is prolonged in an apparent effort to maintain the intracellular calcium dynamics necessary for contraction (Janczewski et al., 2002). Such electrical remodeling is also observed in chronic heart failure, atrial fibrillation and in several genetic and pharmacological models, suggesting that ion channels are under homeostatic regulatory control (Nattel et al., 2007; Rosati and McKinnon, 2004; Schmitt et al., 2014). These observations raise the possibility that the heart monitors its own efficacy and adaptively remodels its electromechanical tone. However, significant implications to the pathophysiology of heart disease notwithstanding, the extended lifetimes and complexity of traditional mammalian models, as well as the necessity of their cardiovascular system for organismal survival, have confounded our ability to examine these potential homeostatic responses.

In seeking to develop a more tractable model for understanding this process, we reasoned that all electromechanical biological oscillators must defend themselves against intrinsic and environmental instability and compensate for natural senescence if they are to maintain their efficacy over time. The first primordial cardiomyocytes likely originated in Bilateria, and the transcription factors tinman (Nkx2.5 in mammals), hand and mef2 represent a conserved cardiogenic program linking the evolution of Protostome and Deuterostome hearts (Bishopric, 2005). Despite having an open circulatory system that reverses the direction of flow periodically (Wasserthal, 2007) and a greatly simplified architecture (Rotstein and Paululat, 2016), Drosophila has proven to be of significant utility in the study of cardiac development and physiology, revealing mechanisms of cardiogenesis and heart function that are highly conserved with mammals (den Hoed et al., 2013; Frasch, 2016; Monnier et al., 2012; Neely et al., 2010). Consequently, Drosophila serves as a uniquely high-throughput ‘pioneer’ genetic model for uncovering conserved pathways involved in cardiomyocyte development and function.

A variety of imaging modalities have investigated the molecular mechanisms underlying cardiac function in Drosophila (Ocorr et al., 2014). Most notably, an in situ preparation has been used to isolate the intrinsic regulators of cardiac performance in a defined physiological solution and in the absence of neuronal input. A complementary approach would be to monitor heart function in intact animals, where the full suite of intrinsic, environmental and homeostatic processes regulating the heart would be accessible to investigation, with minimal decay in cardiac performance for hours after preparation. While simple transmitted or reflected luminance measures can robustly monitor the rhythm of the heart in vivo, one cannot accurately measure heart wall displacement using this approach. Optical coherence tomography has been utilized to measure heart rhythm and displacement in vivo, and has successfully uncovered several novel genes affecting cardiac function (Choma et al., 2006; Wolf et al., 2006; Li et al., 2013; Alex et al., 2015). However, increasing heart wall contrast relative to the lumen and surrounding tissues could yield further improvements in our ability to assess cardiac function in the intact animal.

In this study, we present a high resolution fluorescence imaging and analysis toolset for measuring cardiac function in intact, unanesthetized Drosophila melanogaster. Using this platform, we demonstrate that, as in humans, normal aging manifests primarily as defects in relaxation (diastole) while preserving contractile performance, suggesting a conserved susceptibility to aging-related declines in the electrical, biochemical or structural processes facilitating relaxation. We also uncover a critical role for a heteromeric two-pore potassium channel in maintaining cardiac rhythmicity during aging, which appears dispensable for heart function early in life but is critical for preventing fibrillatory cardiac arrest in aged animals. We propose that the robustness, speed and resolution of this in vivo platform will significantly increase the utility of Drosophila in understanding conserved mechanisms of cardiac aging and homeostasis.

Results

Imaging cardiac performance in intact, unanesthetized Drosophila

We developed a fluorescence-based approach for imaging the heart in intact, unanesthetized Drosophila. Briefly, the red fluorophore tdTomato was transgenically expressed in working cardiomyocytes using a newly discovered heart enhancer R94C02 (Tables 1 and 2). Intact flies were attached to coverslips using ultraviolet-activated optical cement and placed in the optical path (Figure 1A). Excitation light was spatially limited to the region of the dorsal abdomen containing the second and third chambers of the heart using a digital-micromirror projector, with the fluorescence emitting back through the cuticle captured on a sCMOS camera operating at 120 frames per second (Figure 1B).

Figure 1 with 1 supplement see all
Imaging cardiac performance in intact, unanesthetized Drosophila.

(A) Full view of the intact preparation, with imaging region of interest (blue). The head and legs are freely moving, while the wings, dorsal thorax and dorsal abdomen are affixed to the coverglass with optical cement. (B) Electro-optical diagram of the imaging system. (C) Single anterograde heartbeat at half frame-rate with heart wall position (yellow), initiation of contraction (red triangles) and of relaxation (blue triangles) calls. (D) Associated YT kymograph (magenta) with heart wall detection (white dots). (E) Corresponding digitization, segmented into anterograde (white) and retrograde (grey) heartbeat epochs. The triangles denote the initiation (red) and end (blue) of contractions. (F) Two-dimensional probability map of heart chamber diameter and heartbeat duration with median +/− quartile overlay for systole (red) and diastole (blue). (G) Two-dimensional probability map of fractional shortening and systolic interval. All data in this figure are representative and from the 10-day-old w1118 wild-type dataset. See Materials and methods for all functional parameter definitions and their derivation. Scale bars: (black vertical) 75 µm, (blue horizontal) 1 s. See also Video 1.

https://doi.org/10.7554/eLife.20851.002

Using this in vivo preparation and an automated feature detection algorithm based on maximal contrast, we unambiguously tracked the heart wall across the cardiac cycle in intact animals (Figure 1C,D). We then developed a segmentation algorithm that converted this digital representation of chamber diameter over time into discrete contraction and relaxation events (Figure 1C–E, Video 1). This segmentation allowed us to derive a diverse set of heart functional parameters, including estimates of cardiac output and stroke volume. We refer the reader to the Materials and methods section for a detailed explanation of the algorithms and formulas used.

Video 1
Heartbeat visualization, digitization and segmentation.

One-third speed video of the 10 day adult female displayed in Figure 1C–E, with heart wall position calls (yellow) and the attending transformation into heart chamber diameter as a function of time in a 1 s streaming window. The initiation and end of each contraction are specified by a red and blue triangle, respectively. Note the periodic reversal in the direction of heartbeat peristalsis.

https://doi.org/10.7554/eLife.20851.004

In wild-type animals, heartbeats are very consistent and can be visualized en masse by assembling a two-dimensional probability map of chamber diameter across the cardiac cycle (Figure 1F) or fractional shortening versus the systolic interval (Figure 1G), which measures the percentage reduction in chamber diameter during a contraction. Both visualizations demonstrated that the mechanical rhythm of the heart is reproducible between animals. This consistency is also evident in the four primary measurements of the cardiac cycle: the systolic interval, diastolic interval, end systolic diameter, and end diastolic diameter. The standard deviations for these four measurements are between 7% and 11% for our wild-type dataset of young animals between 10 and 30 days of age (n.s., Kruskal-Wallis one-way ANOVA followed by Dunn’s multiple comparisons test, n = 68), demonstrating that this in vivo preparation is highly reproducible.

We next performed a series of control experiments to assess the stability of the preparation. The excitation light intensity utilized did not appreciably alter the heartbeat waveform but at higher light intensities, the kinetics of contraction and relaxation accelerated significantly (Figure 1—figure supplement 1A,B, R2 = 0.93). Furthermore, the preparation was stable for at least two hours, with cardiac output maintained even after 19 hr in flies kept hydrated overnight (Figure 1—figure supplement 1C,D), establishing that our approach is not subject to meaningful variability associated with decay in the health of the preparation.

Normal aging manifests as diastolic dysfunction while preserving contractile performance

While modifiable risk factors including elevated blood pressure, tobacco use, abnormal blood sugar levels, physical inactivity and obesity strongly exacerbate the incidence of heart disease (Yancy et al., 2013), aging-related changes in the structure and physiology of the heart also influence disease progression (Lakatta et al., 2014). Increases in arterial load, ventricular hypertrophy and diastolic dysfunction are well established in healthy aging hearts (Carrick-Ranson et al., 2012; Cheng et al., 2009; Lakatta et al., 2014). At the molecular level, animal models have revealed clear alterations in action potential duration and calcium handling with age (Feridooni et al., 2015). However, the relative contribution of physiological remodeling in cardiomyocytes to declines in cardiac performance has proven difficult to deconvolve from defects arising from arterial overload and structural hypertrophy.

To establish whether our Drosophila platform might represent a simplified model to study cardiomyocyte senescence in vivo, we aged female flies to determine how heart function would change over time. Previous research using an in situ dissected preparation in Drosophila has demonstrated progressive declines in cardiac rhythmicity during the first five weeks of life (Ocorr et al., 2007). In our in vivo preparation, the heart rate and cardiac performance of flies was remarkably stable for up to 30 days of age in two ‘wild-type’ genetic backgrounds, w1118 and Canton S, which we subsequently grouped for further analysis (6.9% maximal variance in heart rate between these two groups at 10, 20 and 30 days of age, n.s., Kruskal-Wallis one-way ANOVA followed by Dunn’s multiple comparisons test). At 50 days of age, flies displayed a small decrease in heart rate, (Figure 2A, R2 = 0.84) which primarily reflected a prolongation of the systolic interval (Figure 2—figure supplement 1A). Only after 50 days of age did the diastolic interval lengthen (Figure 2—figure supplement 1B) and cardiac output per second decline (Figure 2B).

Figure 2 with 1 supplement see all
Normal aging is characterized by a diastolic decline with preserved contractility.

Various cardiac functional parameters presented by age in a combined w1118 and Canton S dataset, n = 18 to 30 animals per time-point: (A) Median heartbeat with heart rate (inset). (B) Estimated cardiac output per second (blue) and per stroke (red). (C) Median heart wall velocity with peak velocities of contraction (red dots) and relaxation (blue dots). (D) Probability histograms of the time from initiation of contraction to peak contraction velocity (red) and from the peak contraction velocity to peak relaxation velocity (blue). The shaded areas in panel B represent the mean +/− s.d., with regressions plotted as dotted lines.

https://doi.org/10.7554/eLife.20851.005

Although we observed significant reductions in total cardiac output in aged animals (Figure 2B), the mechanical performance of each individual contraction was remarkably preserved. We observed no significant changes in stroke volume (Figure 2B, red), chamber diameter (Figure 2—figure supplement 1C), or fractional shortening (Figure 2—figure supplement 1D), suggesting that the observed decline in cardiac performance might reside in the temporal rather than spatial domain. Indeed, the most striking change associated with aging was an increased latency in transitioning from systole to diastole, which primarily reflected a decrease in the velocity of relaxation (positive values in Figure 2C) and an increase in the time period from peak contraction velocity to peak relaxation velocity (Figure 2D, blue, R2 = 0.73). In contrast, the kinetics of contraction was much less affected (Figure 2C,D). Similar diastolic decline with preserved contractile performance accounts for an increasing fraction of heart failure cases in humans (Borlaug, 2014; Sharma and Kass, 2014), suggesting a conserved susceptibility to aging-related declines in the structural or biochemical processes facilitating relaxation.

A pair of K2P subunits, sandman and galene, are required for terminating systole in aged animals

The rhythmic contraction and relaxation of the heart requires a precisely tuned series of ionic conductances that entrain the influx and efflux of calcium across the sarcolemnal and sarcoplasmic reticular membranes (Bers, 2008; Monfredi et al., 2013). In cardiomyocytes, outward potassium currents mediating repolarization are essential for terminating systole and suppressing dysrhythmic afterdepolarizations (Nerbonne and Kass, 2005). One physiological hallmark of failing hearts is the progressive loss of these repolarizing currents (Beuckelmann et al., 1993). We therefore investigated the repolarizing mechanisms maintaining diastolic function and normal rhythm in Drosophila. Previous work has implicated a number of potassium channels important in the repolarization to, or maintenance of, the cardiac resting potential in Drosophila, including KCNQ (Ocorr et al., 2007) and the K2P channel ORK1 (Lalevée et al., 2006).

While several voltage-gated potassium channels have well established roles in cardiac repolarization, a number of K2P channel family members are expressed in the human heart but their physiological relevance remains an active area of investigation (Schmitt et al., 2014). TASK-1 (K2P3.1) has been implicated in chronic atrial fibrillation, where TASK-1 protein levels are increased and action potential duration is shortened in a TASK-1 dependent manner relative to controls (Schmidt et al., 2015). In the larval heart of Drosophila, the K2P family member ORK1 appears to fine tune the rate of slow diastolic depolarization (Lalevée et al., 2006). Transcriptional profiling has previously revealed a putative two-pore potassium channel subunit, CG8713, recently named sandman (Pimentel et al., 2016), with enriched mRNA expression in the Drosophila heart relative to other tissues (Robinson et al., 2013). Reasoning that this gene may play a role in cardiac repolarization, we confirmed its expression in the fly heart using RT-PCR and generated a small deletion of CG8713 and the adjacent gene CG8712 (Figure 3—figure supplement 1A,C). At 50 days of age, the hearts of sandman mutants displayed a marked inability to transition from heart contraction (systole) to heart relaxation (diastole), with some animals rarely or never displaying heart relaxation (Figure 3A, Video 2).

Figure 3 with 2 supplements see all
Diastolic failure in sandman mutants.

(A) Representative YT kymographs of 50-day-old animals. Scale bars: (black vertical) 75 µm, (blue horizontal) 1 s. (B) Median heartbeat per genotype at 50 days of age. ⊕, clean excision of the mutagenic piggyBac insertion e00867. (C) Two-dimensional probability map of fractional shortening and systolic interval at 50 days of age. See also Video 2.

https://doi.org/10.7554/eLife.20851.008
Video 2
50-day wild-type and sandman heartbeat visualization.

One-third speed videos of 50-day adult wild-type (upper video) and sandman (lower two videos) females, with heart wall position calls (yellow) and the attending transformation into heart chamber diameter as a function of time in a 1 s streaming window. The initiation and end of each contraction are specified by a red and blue triangle, respectively.

https://doi.org/10.7554/eLife.20851.012

This phenotype appears to be heart autonomous and specifically due to the loss of sandman, not CG8712. Expression of the sandman cDNA in cardiomyocytes using R94C02::Gal4 rescued the functional defects observed, including the median heartbeat waveform (Figure 3B) and the probability map of fractional shortening versus systolic interval (Figure 3C). We next screened other putative K2P channel subunits and uncovered a similar heart-autonomous role the heart expressed K2P subunit CG9194 (henceforth galene) (Figure 3—figure supplement 1B,C), for which knockdown also led to a marked decline in cardiac function. Heterozygous animals expressing dsRNA selectively targeting galene in the heart using tinCΔ4::Gal4 displayed a dispersion in fractional shortening versus systolic interval (Figure 3—figure supplement 2A), a significant reduction in cardiac output per second and per stroke (Figure 3—figure supplement 2B), and a prolongation of the systolic interval relative to controls (Figure 3—figure supplement 2C). As in sandman mutants, this defect primarily reflects a difficulty in transitioning from systole to diastole; the median heartbeat waveform exhibited a clear reduction of diastolic function relative to controls (Figure 3—figure supplement 2D).

The sandman phenotype displays an age-dependent progression

Defects in cardiac repolarization can manifest as life threatening arrhythmias, but their pathophysiology is complex due to compensatory repolarization reserve and arrhythmogenic electrical remodeling (Nattel et al., 2007). For example, atrial fibrillation exhibits an age-dependent penetrance that reflects self-reinforcing electrophysiological remodeling and normal aging. The action potential is shortened, which facilitates reentrant excitation, causing further shortening of the action potential thereby completing the feed-forward loop. Conversely, aging hearts exhibit action potential prolongation, which augments contractility but also predisposes the heart to Torsades des Pointes tachyarrhythmia, a ventricular rhythm defect that can lead to sudden cardiac death. Such age-dependent pathogenesis is evident in a Drosophila model of long QT syndrome where KCNQ potassium channel mutants develop progressive dysrhythmia (Ocorr et al., 2007).

We systematically quantified the age-dependent progression of diastolic failure in sandman mutants. sandman mutants displayed only minor defects in cardiac function early in life. Cardiac output per second and per stroke were statistically indistinguishable from wild-type at 10 days of age (Figure 4A,B), with only modest increases in the systolic interval (Figure 4C). However, by 30 days of age heart function was severely compromised; cardiac output per second and per stroke declined precipitously (Figure 4A,B) while the systolic and diastolic durations increased dramatically (Figure 4C,D). By 50 days of age, median total cardiac output per second had declined approximately 100-fold (Figure 4A). This progression can also be readily observed in two-dimensional probability maps of chamber diameter and fractional shortening (Figure 4—figure supplement 1) and can be rescued by expressing sandman cDNA selectively in cardiomyocytes of sandman mutants (Figure 4A–D, green boxplots) but not when expressed selectively in body wall muscles excluding the heart (see Figure 4—figure supplements 2 and 3 for detailed statistics for all measurements by age and genotype).

Figure 4 with 3 supplements see all
Progressive heart failure in sandman mutants.

(A–D) Estimated cardiac output per second (A) and per stroke (B) were well fit by a Boltzmann sigmoidal regression and the systolic (C) and diastolic (D) intervals were well fit by single exponential growth regression curves for wild-type (grey), sandman (red), cardiomyocyte rescue of sandman using R94C02::Gal4 (green) and the clean excision (pink) at specified ages. n = 7 to 27 animals per genotype and age. The shaded areas represent the mean +/− s.d., with the regressions plotted as dashed lines.

https://doi.org/10.7554/eLife.20851.013

Sandman and galene jointly encode a heteromeric potassium channel in vitro

Although historically considered background ‘leak’ currents that influence the resting membrane potential of cells, K2P channels are gated by diverse physiochemical stimuli and may therefore couple the activity of tissues to their environment (Enyedi et al., 2010). sandman was recently implicated as a dopamine-induced potassium current that operates as part of a homeostatic sleep switch (Pimentel et al., 2016). We characterized the electrophysiological behavior of sandman and galene by expressing them in Chinese hamster ovary (CHO) cells. Consistent with their similar phenotypes in vivo, sandman and galene most likely encode two subunits that form a heteromeric ion channel in vitro. Neither subunit expressed alone was sufficient to give rise to an appreciable conductance in CHO cells whereas co-expression resulted in large outward currents at depolarized potentials (Figure 5A). This conductance is highly selective for potassium and, in contrast to the open rectifier ORK1 (Goldstein et al., 1996), displays outward rectification that is only partially due to block by divalent cations (Figure 5B), a conserved feature of many K2P channel subtypes (Schewe et al., 2016). Importantly, this functional co-assembly appears selective; the two most closely related K2P subunits, CG1688 and CG10864, do not form functional heteromers with either Sandman or Galene (Figure 5—figure supplement 1). Heteromultimerization of potassium channel subunits is a well-established mechanism for increasing electrophysiological diversity of tetrameric potassium channels as well as dimeric K2P channels (Yang and Nerbonne, 2016). Considering that several K2P channel subtypes are functionally silent as homodimers in vitro (Enyedi et al., 2010; Goldstein et al., 2005), heteromeric complementation, as observed between Sandman and Galene (Figure 5A), may be of considerable significance to the physiology of this family.

Figure 5 with 1 supplement see all
sandman and galene jointly encode a potassium channel.

(A) Representative whole-cell currents in physiological K+ and Na+ gradients from Sandman (n = 5), Galene (n = 6), and co-transfection of both (n = 11) during voltage steps (below). (B) Normalized whole-cell currents from voltage ramps in various bath solutions. The dotted line plots the I/V curve for a hypothetical ion channel with no rectification in symmetric K+. The inset plots the observed reversal potential compared to a potassium-selective conductance (dashed line) at various [K+] in/out ratios. The internal pipet solution is (in mM) 150 K+, 5 Na+, 3 Mg2+, 161 Cl, 10 HEPES, pH 7.4 (in mM). The bath solution [K+] and [Na+] or [NMDG+] are as indicated (in mM), excepting the ‘Divalent-free’ solution which substitutes 2 mM EDTA for the divalent cations. n = 9 cells. All pooled data represent the mean +/− s.d. All voltage potentials are relative to ground.

https://doi.org/10.7554/eLife.20851.017

The sandman phenotype likely reflects a loss of repolarization rather than structurally congestive remodeling

To our knowledge, Drosophila represents the only model organism where the pumping action of the heart is acutely dispensable for adult survival. We therefore characterized the terminal phenotype of sandman mutants to determine whether it may reflect congestive structural remodeling or a heart that is constitutively contracting. To differentiate these possibilities, we acutely injected the calcium chelator EGTA into the abdominal cavity of intact 110 days old sandman animals and compared the heart’s contractile state before and immediately after. EGTA robustly terminated the persistent contractions observed in sandman mutants, establishing that cardiomyocytes were actively contracting in a Ca2+ dependent manner rather than locked into congestive cytoskeletal remodeling (Figure 6A). Similarly, acute injections of the potassium ionophore valinomycin also terminated the prolonged contractions observed in sandman mutants (Figure 6A), confirming that the phenotype was dependent on the intracellular concentration of potassium ions and likely reflected a failure to fully repolarize.

In vivo pharmacology and sarcomere dynamics implicate dyssynchronous and regenerative Ca2+ in maintaining persistent systole.

(A) Representative heart kymographs from 110-day-old sandman males before and acutely after intra-abdominal injection of the Ca2+ chelator EGTA or the potassium ionophore valinomycin. n = 3. (B) Micrograph of a dissected adult Drosophila expressing a GFP trap of the z-line protein α-actinin [CC01961]. (C–D), Representative kymographs (magenta) of second chamber right and left cardiomyocyte sarcomere dynamics from intact 30-day-old animals, as visualized intravitally using the α-actinin GFP-trap. Automated detection of one z-line for each cardiomyocyte (green/white), quantified as relative position over time (upper trace, left cardiomyocyte signal inverted), with net coherence between z-lines (middle trace) and integral coherence (bottom trace). Scale bars: (black vertical) 10 µm, (blue horizontal) 1 s. n = 9 for wild-type and four for sandman. See also Video 3.

https://doi.org/10.7554/eLife.20851.020

To further characterize the constitutively contracted state of aged sandman hearts, we directly imaged sarcomere dynamics in vivo using a GFP protein trap of the z-line protein α-actinin (Figure 6B). We quantified the relative coherence of sarcomeres between left and right cardiomyocyte pairs by calculating the fraction of time in which they were moving in unison towards or away from the midline, which reflects systole and diastole respectively. As expected, wild-type sarcomeres contract and relax in unison (Figure 6C, Video 3). In sandman mutants, individual sarcomeres exhibited a fibrillatory state during extended systoles, contracting and relaxing out of phase with one another (Figure 6D, Video 3), suggesting that sufficient ATP was available locally to drive myosin activity, myosin dissociation and the various pumps that can sequester or extrude calcium. Because the contractile state of individual sarcomeres is intimately linked to local calcium cycling (Bers, 2008; Hohendanner et al., 2013; Venetucci et al., 2008), it is possible that the observed dyssynchrony of sarcomere dynamics reflects asynchronous local calcium rise and sequestration in the absence of a coherently cycling action potential.

Video 3
30 day wild-type and sandman sarcomere dynamics.

Real-time videos of 30-day adult wild-type (upper video) and sandman (lower video) females, visualizing the z-lines of the dorsal aspect of the cardiomyocyte pair just posterior to the ostial valves of the second chamber using a protein trap of α-actinin [CC01961].

https://doi.org/10.7554/eLife.20851.021

Discussion

The Drosophila heart represents a reduced system for understanding fundamental mechanisms of heart function and aging

In this study, we establish a genetically accessible in vivo model for understanding the molecular mechanisms regulating cardiac performance during normal aging. Our imaging methodology permits the extended imaging of heart function in intact, unanesthetized animals for several hours, without measurable declines in cardiac performance (Figure 1—figure supplement 1C,D). The resolution and reproducibility of our measurements revealed specific changes to the heartbeat waveform as animals age. Using a total of 146 wild-type flies from six different ages, we visualized approximately 65,000 heartbeats, reconstructing median heartbeat waveforms for each age (Figure 2A) and demonstrating a progressive decline in the kinetics of relaxation but not contraction (Figure 2C,D).

The throughput of this in vivo assay facilitates the identification of novel genes that establish and maintain cardiac function and can therefore complement more complex vertebrate models. In a small scale screen, we identified two K2P channel subunit genes, sandman and galene, which together give rise to a heteromeric potassium channel that appears essential for terminating systole and promoting relaxation in aged animals. Our analysis of sandman mutants revealed that the pumping function of the heart is acutely dispensable for adult Drosophila survival under laboratory conditions (Figure 3A, Video 2). Although cardiac function is not required for zebrafish embryogenesis or the first week of development, heart function is otherwise essential for the survival of all adult vertebrates (Staudt and Stainier, 2012). Differences in heart dispensability likely originate in the decoupling of gas exchange and heart function in larval zebrafish or insects, where sufficient gas exchange can occur via local diffusion. This opens a unique window for understanding the physiological transition to, and maintenance of, fibrillatory arrest without the confounds of organismal or cardiomyocyte death. The future development of in vivo approaches for monitoring cytosolic and sarcoplasmic reticular Ca2+ levels in the beating heart will facilitate a clearer mechanistic view of the biophysical mechanisms initiating and sustaining fibrillatory arrest.

Sandman is required to maintain diverse biological oscillators

Oscillatory behaviors exist across diverse timescales, from migratory cycles in birds to ultrafast spiking neurons in the auditory system. One conserved feature of oscillators is their capacity to be modulated by internal and external cues, thus adapting the system to immediate physiological needs or entraining the phase of the cycle to the external world. Interestingly, sandman appears to play a central role in two radically different biological oscillations: organismal sleep-wakefulness (Pimentel et al., 2016) and the contraction-relaxation of aging hearts (this paper). Pimental and colleagues demonstrated that Sandman-dependent potassium currents are upregulated by dopamine via a G-protein cascade that is pertussis toxin sensitive. This increase in potassium ‘leak’ significantly reduces the excitability of sleep-promoting dFB neurons and therefore tips the balance of the cycle towards wakefulness. Phenotypically, our results suggest that sandman plays a critical role in maintaining a contraction-relaxation oscillator during aging. Loss of sandman does not grossly perturb cardiac rhythmicity in young animals but the balance between contraction and relaxation degenerates catastrophically with age, leaving the heart in a persistently contracted state.

While the molecular mechanism by which pertussis-toxin sensitive G-protein signaling upregulates Sandman currents is not yet understood, a variety of downstream signaling events are known to affect the activity and localization of ion channels (Inanobe and Kurachi, 2014). Similar pathways play a significant and complex role in the neurohormonal regulation of the mammalian heart, for instance by the counterbalanced G-protein coupled pathways activated by the sympathetic and parasympathetic nervous systems (Mangoni and Nargeot, 2008). Aging human hearts exhibit diminished responsiveness to these modulatory pathways, suggesting senescent defects in signal transduction or a system driven to the limits of its dynamic range (Kaye and Esler, 2008). Similarly, work in Drosophila has demonstrated that the heart is sensitive to neural input (Dulcis and Levine, 2005). It will be interesting to determine whether extrinsic or intrinsic regulatory signals tune Sandman activity during aging and thus optimize the heart’s contraction-relaxation balance in a fashion analogous to that observed in the sleep center of the brain.

Diastolic decline with preserved contractility may represent a conserved feature of aging cardiomyocytes

Young hearts possess abundant repolarizing reserve and can often compensate for losses in individual conductances without impeding the heart’s ability to rapidly relax (Nattel et al., 2007). For reasons that are not well understood, this reserve steadily declines with age in humans and animal models, leaving the heart susceptible to late-onset dysrhythmias. The cardiac phenotype observed upon loss of sandman is strikingly age-dependent, transitioning from grossly normal heartbeat in young animals to complete fibrillatory arrest later in life. Our in vivo pharmacological results implicate defects in action potential repolarization, consistent with the rectification behavior of the Sandman/Galene heteromeric channel (Figure 5B). These observations suggest that a progressive loss of repolarization reserve may represent a conserved feature of cardiac aging. However, the pathophysiological mechanisms underlying this age-dependent loss are not known.

In humans, an increasing number of heart failure cases display significant defects in relaxation while largely preserving contractile performance, termed 'Heart Failure with preserved Ejection Fraction' (HFpEF) (Borlaug, 2014; Sharma and Kass, 2014). Owing to the closed nature of the vertebrate cardiovascular system, the pathophysiology of such diastolic decline reflects a diverse and complicated assemblage of causal mechanisms including cardiomyocyte dysfunction, structural remodeling and increased vascular resistance. Despite dramatic differences in the architecture of their cardiovascular systems, aging Drosophila similarly exhibit preferential diastolic decline (Figure 2). Although many pathways likely contribute to the pathophysiology of HFpEF, our results suggest that cardiomyocytes across phylogeny may possess a conserved active mechanism driving this differential outcome.

Previous work has uncovered several potential mechanisms that may differentially regulate systolic and diastolic tone. Mechanically, a growing body of literature has implicated isoform switching and postranslational modifications of the giant macromolecular spring Titin (Linke and Hamdani, 2014). Patients experiencing heart failure with preserved ejection fraction exhibit Titin hypophosphorylation, which increases the resting tension of cardiomyocytes and impairs diastolic function (Hamdani et al., 2013). Conversely, increasing Titin compliance experimentally has a beneficial effect on diastolic performance but compromises elements of systolic function, notably the Frank-Starling reflex (Methawasin et al., 2014). Electrophysiologically, there is considerable evidence that action potential duration is differentially remodeled in aging hearts, heart failure and atrial fibrillation (Beuckelmann et al., 1993; Heijman et al., 2014; Janczewski et al., 2002), which may ameliorate the initial dysfunction but predisposes the heart to later dysrhythmia. In animal models and patients exhibiting chronic heart failure, several potassium channel subunits appear downregulated (Beuckelmann et al., 1993; Nattel et al., 2007), prolonging the action potential in an apparent attempt to augment contractile function. Such decline has also been observed in aging Drosophila, where KCNQ transcripts appear downregulated and animals lacking KCNQ display progressive dysrhythmia in situ (Ocorr et al., 2007).

Together, these observations suggest that contractile function may be adaptively regulated by mechanisms that are conserved across phylogeny and that age-related diastolic decline and increased susceptibility to dysrhythmia may represent unintended side effects of this compensation. Our results also implicate a heteromeric potassium channel as a critical effector for maintaining normal rhythmicity during aging, suggesting that age-dependent cardiomyocyte membrane properties may play a key role in maintaining cardiac function into old age. Several important questions remain unaddressed. How does the heart or the brain sense, integrate and respond to alterations in contractile efficacy? Do these mechanisms exacerbate declines in diastolic function during normal aging and to what extent do these homeostatic mechanisms contribute to the pathogenesis of heart disease? The acute dispensability of the fly heart pumping, the extensive genetic tools available in Drosophila, and the intravital imaging system we developed provide an exciting opportunity for exploring this dynamic nexus between cardiac physiology, aging and disease.

Materials and methods

DNA constructs

DNA plasmids used for fly transgenesis and heterologous cell transfection were assembled using standard molecular biological techniques and sequenced to confirm accuracy and identity. Sub-cloning details and plasmid descriptions are presented in Table 1.

Table 1

DNA constructs.

https://doi.org/10.7554/eLife.20851.022
Plasmid IDPlasmid nameInsert source5’ Primer3’ PrimerDestination VectorRestriction SubcloningComments
pMK110xUAS-IVS-Syn21-tdTomato-p10pDEST HemmarR (Addgene #31222)ataaggtaccAACTTAAAAAAAAAAATCAAAATGGTGAGCAAGGGCGAGatattctagaTTACTTGTACAGCTCGTCCATGCCpJFRC81 (Addgene #36432)KpnI - XbaIIntermediate plasmid - for fly transgenesis (Han et al., 2011; Pfeiffer et al., 2012)
pMK3R94C02::tdTomatoJanelia Farms, amplified from Drosophila genometactagtACTTTTCCGCGCCGTCTGatatgctagcGGAAACAGACGCAAAGACTGACpMK1 (this paper)HindIII - NheICardiomyocyte enhancer expressing tdTomato for fly transgenesis - 5’ primer was phosphorylated to facilitate blunt ligation using Klenow fragment
pMK17pAc5.1B_GFP-CG8713BDGP cDNA RE21922 in clone #UFO03925aaaagcggccgcATGTCCTCCCGACGCatattctagaTTAGGAGGTGCGGCACpAc5.1B_GFP (Addgene #21181)NotI - XbaIIntermediate plasmid - for S2 cell expression (Stapleton et al., 2002)
pMK18pAc5.1B_GFP-CG10864PCR from Drosophila genome (single exon)aaaagcggccgcATGGCCAGCAAATTTCAGAGatattctagaCTAGTAGTAATCATCCTCGTACpAc5.1B_GFP (Addgene #21181)NotI - XbaIIntermediate plasmid - for S2 cell expression
pMK21pAc5.1B_GFP-CG1688BDGP cDNA GH04802 in clone #UFO05944aaaagcggccgcATGTCCGACGTTGAGCAGatattctagaTTATCCATCCGCGCGGpAc5.1B_GFP (Addgene #21181)NotI - XbaIIntermediate plasmid - for S2 cell expression
pMK22pACU_GFP-CG8713pAc5.1B_GFP- CG8713 (this paper)KpnI - ApaI insert from source vectorKpnI - ApaI insert from source vectorpACU (Addgene #58373)KpnI - ApaIUAS::GFP-CG8713 rescue construct - for fly transgenesis
pMK23peGFP_C1-CG8713pAc5.1B_GFP- CG8713 (this paper)EcoRI - ApaI insert from source vectorEcoRI - ApaI insert from source vectorpeGFP_C1 (Clontech)EcoRI - ApaIFor expression in mammalian heterologous cells
pMK24pAC5.1B_eGFP-CG9194BDGP cDNA FI03418 in clone #UFO11253aaaaagcggccgcATGTCGGGTAGGCGGGCCCAgcagcctctagaCTAATCCTCATCCTGCTCGTCGTCATCATCCpAc5.1B_GFP (Addgene #21181)NotI - XbaIIntermediate plasmid - for S2 cell expression
pMK25peGFP_C1-CG9194pAC5.1B_eGFP- CG9194 (this paper)EcoR1 - SacII insert from source vectorEcoR1 - SacII insert from source vectorpeGFP_C1 (Clontech)EcoR1 - SacIIFor expression in mammalian heterologous cells
pMK29peGFP_C1-CG1688pAC5.1B_eGFP- CG1688 (this paper)tataGCTAGCggtaccaacatggtgagcaaggtatacgggccctctagaTTATCCATCCpeGFP_C1 (Clontech)NheI - ApaIFor expression in mammalian heterologous cells
pMK30peGFP_C1- CG10864pAC5.1B_eGFP- CG10864 (this paper)EcoR1 - ApaI insert from source vectorEcoR1 - ApaI insert from source vectorpeGFP_C1 (Clontech)EcoR1 - ApaIFor expression in mammalian heterologous cells

RT-PCR

Fifty fly hearts were microdissected from the adult abdomen of mixed age w1118 flies and total RNA was isolated using the ZR RNA MicroPrep Kit (Zymo Research, Irvine, CA). cDNA transcripts were generated using Superscript III RT and oligo dT primers (Thermo Fisher) and PCR amplification was tested using GoTaq Green Master Mix (Promega) using the following primers:

sandman 5’ TACAGAGCGCGCAAACATA 3’ AGGATTTCCGGCTACCTATCG

galene 5’ TTTGTGGCTCGTACGGATCG 3’ CTAATTTGCCGCTCGGTTGG

Drosophila genetics and husbandry

All chromosomal aberrations and transgenic insertions used in this study are detailed in Table 2. Transgenic elements generated in the course of this study were inserted into specific attP docking sites within the Drosophila genome using phiC31-mediated integration (Bischof et al., 2007). The deletion of sandman, e00867-e00152, was generated using FLP-FRT mediated recombination of two piggyBac elements in trans and confirmed using PCR (Parks et al., 2004; Thibault et al., 2004). Clean excision of the piggyBac elements was performed as previously described (Parks et al., 2004; Thibault et al., 2004). Animals were raised on standard cornmeal, yeast, agar, molasses formula and kept in a diurnal 12 hr light: 12 hr dark 70% humidity-controlled incubator (Darwin Chambers, St. Louis, MO). All experimental adult flies were raised at 20°C excepting RNAi knockdown, which was performed at 25°C to increase dsRNA expression. For aging, 20 female and 10 male flies were transferred to fresh unyeasted vials every 7 days. Approximately 18 hr before imaging, a wedge of rinsed and water saturated cellulose acetate (Genesee Scientific, San Diego, CA) was added to the fly vials to ensure adequate hydration of the flies. CG9194 was named galene after the ancient Greek goddess of calm seas.

Table 2

Drosophila genomic aberrations and transgenic insertions.

https://doi.org/10.7554/eLife.20851.023
Chromosomal ElementLocationSourceDescription
R94C02::tdTomatoChr. II (attP40)This paperR94C02 enhancer expressing tdTomato in cardiomyocytes and a subset of other muscles within the adult fly. Integrated into the attP40 docking site. We used this transgenic for all heart wall imaging experiments.
PBac{RB}CG8713 [e00867]Chr. IIExelixis via BloomingtonpiggyBac{RB} insertion proximal to the 1st splice acceptor of the CG8713 mRNA. This insertion has a strong heart function phenotype.
PBac{RB}CG8712 [e00152]Chr. IIExelixis via BloomingtonpiggyBac{RB} insertion into the CG8712 coding sequence. This insertion does not have a discernible heart phenotype.
PBac{RB}CG8713_CG8712 [e00867_e00152 deletion, mW+]Chr. IIThis paperFLP-FRT mediated deletion of the genomic DNA between e00867 and e00152, which comprises the entire CG8713 coding sequence and most of the CG8712 coding sequence. RB(+) to RB(+) recombination reconstitutes a single PBac{RB} after the intervening sequence is deleted.
[e00867_e00152 deletion]Chr. IIThis paperClean excision of PBac{RB}CG8713_CG8712 [e00867_e00152 deletion, mW+]. This chromosome harbors the deletion without any residual pBac[RB] sequence.
CyO, P{Tub-PBac\T}Chr. IIBloomingtonSource of piggyBac transposase activity for generating clean excisions, which were identified by the loss of mini-white eye pigmentation.
e00867 [clean excision]Chr. IIThis paperClean excision of PBac{RB}CG8713 [e00867] which reverts the observed heart phenotype.
UAS::GFP-CG8713Chr. III (vk00005)This papernsertion of plasmid pMK22 into the attP docking site vk00005. Used to demonstrate cardiomyocyte rescue of CG8713 (anubis).
R94C02::Gal4Chr. III (attP2)BloomingtonIn a screen for new heart-specific Gal4s, we found R94C02::Gal4 as a complement to tin.CΔ4::Gal4. It is expressed in cardiomyocytes and a subset of other muscles. (Pfeiffer et al., 2008)
tin.CΔ4::Gal4Chr. IIIManfred Fraschtin.CΔ4 is expressed in the heart and a subset of other muscles within the fly. (Lo and Frasch, 2001)
R37B05::Gal4Chr. III (attP2)BloomingtonIn a screen for new heart-specific Gal4s, we found R37B05::Gal4 to be expressed in bodywall muslces but not the heart.
Df(2R)BSC267Chr. IIBloomingtonA molecularly defined deletion spanning the CG8713 (sandman) locus.
Df(3L)BSC431Chr. IIIBloomingtonA molecularly defined deletion spanning the CG9194 (galene) locus.
P{KK110628}VIE-260B Chr. IIVDRC #v108758UAS::CG9194 dsRNA for tissue-specific knockdown of galene. UP-TORR does not predict any off-target effects.
P{w[+mC]=PTT-GC Actn [CC01961]Chr. XBloomington, this paperGFP translational fusion of α-Actinin, which is localized to the z-lines of all muscles. We recombined away the yellow and white alleles.
Canton SN/AJan LabWild-type stock
w [1118]Chr. XJan LabWild-type stock, outcrossed to Canton S 6x
M{vas-int.Dm}ZH-2A; +; PBac{y[+]-attP-9A}VK00005Chr. X and IIIBloomingtonUsed for phiC31 mediated integration.
P{nos-phiC31\int.NLS}X; P{CaryP}attP40Chr. X and IIBloomingtonUsed for phiC31 mediated integration.

Intravital imaging

We used a modified Olympus BX51WI microscope for all video acquisition. Excitation light was provided by a DMD projector (DS + 6K-M, Christie Digital Systems, Cypress, CA) with the green light excitation intensity (538–568 nm, 4.3 mW/mm2) and spatial extent controlled by PsychoPy (Peirce, 2007), using a standard 8 bit RGB tiff file as the signal. The projector was coupled to the microscope using a relay formed by two 150 mm focal length achromatic doublet lenses (Thorlabs Inc., Newton, NJ), placed into a tube assembly and attached to a second camera port above the vertical illuminator (Olympus U-DP and U-DP1XC), with the filter cube installed in the U-DP. We utilized a 20x/1.0 NA water-immersion objective (Olympus) and a sCMOS camera (PCO-Tech Inc., Romulus, MI), triggered using the vertical-sync of the projector signal as frequency doubled by a Master-8 stimulator (AMPI Jerusalem, Israel). Each fly was briefly anesthetized using ice, coupled to a No. one coverslip using Norland Optical adhesive #61 cured using a 365 nm 3 watt UV LED (LED Engin Knc., San Jose, CA) for 10 s and allowed to recover for 10 min before imaging. The coverslip was mounted to the microscope using an assembly containing a small goniometer (Thorlabs GN-05) which allowed the pitch of each fly to be optimized. The optical path is as follows: water immersion objective, coverslip, optical cement, and intact fly. Images were acquired in global shutter mode using a 255 MHz clock with a 6 ms exposure at 120 frames per second. Each animal was recorded for 90 s, with 10,800 frames written directly to an array of 15 spindle disks using a RAID controller with write back cache enabled (LSI SAS9280, Avago Technologies, San Jose, CA). Acquisitions were Gaussian downsampled to 1 µm per pixel and converted to 8 bit using an ImageJ script prior to analysis. All heart wall videos in the manuscript utilized the same transgenic heart marker, R94C02::tdTomato[attP40], as heterozygotes.

Heartbeat digitization

Using a manually-assisted ImageJ script, YT orthogonal kymographs of the heart wall just anterior to the ostial valve of the second heart chamber were generated. At this time, we also manually traced the systolic lumenal area of the second heart chamber, as specified by an average projection of the entire 10,800 frame stack. This assisted script calculated the area of the second chamber in systole and its length, which were appended to the image as metadata for later use. A second ImageJ script detected the right and left heart wall positions in the kymograph independently using a maximal contrast algorithm, refined by a four pixel maximum intensity search medial to that call. These calls were subsequently low pass filtered using an 8-pole 12 Hz Bessel filter (filter poles calculated using online software at http://www-users.cs.york.ac.uk/~fisher/mkfilter/trad.html). Rapid transient displacements resulting from errors in the heart wall detection algorithm were recursively smoothened using preceding and subsequent heart wall position calls. Analysis for the knockdown of galene was performed single-blind, but all other experiments were not blinded to genotype.

Heartbeat segmentation

We automatically segmented each heartbeat into discrete contraction and relaxation events and analyzed these events in detail using VisualBasic scripts written for Diadem 2011 (National Instruments, Austin, TX). The following represents a summary of the material algorithms used to generate the functional parameters presented herein. First, we detected all contraction and relaxation events that fulfilled minimum velocity (75 µm/s), duration (24 ms) and displacement (2.5 µm) criteria. We then eliminated all prospective events where the two heart walls were not moving in unison, which resolved the residual fraction of false heart wall position calls. We next eliminated excess contraction and relaxation calls using nested timing and amplitude criteria so that the principal contractions and relaxations are interleaved 1:1. The initiation and end of contractions were then calculated by walking back to the last zero velocity time point before each contraction and relaxation. The software then acquired the heart diameters when contraction initiated and ended, which represent the end diastolic diameter (EDD) and end systolic diameter (ESD), respectively. These heartbeats were subsequently refined by consolidating compound contractions or relaxations not separated by a minimum duration (24 ms), below the minimum fractional shortening (0.04) or not exhibiting sufficient coherence between the two heart walls.

Heartbeat analysis

A variety of functional parameters were calculated from the segmented heartbeat waveforms and are italicized and underlined for ease of reference. The end diastolic diameter (EDD) and end systolic diameter (ESD) were calculated as described in the previous section. We also quantified heart chamber diameter across the cardiac cycle, denoted by ∀. The systolic interval is the duration of contraction. The diastolic interval is the time elapsed between the end of contraction and the initiation of the next contraction. We observe only the occasional pause at the end diastolic diameter in Drosophila. The fractional shortening is the percentage change in diameter for each contraction where FS = (EDD - ESD)/EDD. The stroke volume of each heartbeat was estimated by modeling the heart as a radially contracting cylinder, using the average systolic diameter and fixed chamber length, in microns, generated for each animal during digitization as fixed variables. The average chamber systolic diameter was divided by the mean ESD to scale the measured ESDs onto the heart model. The estimated chamber systolic diameter for each heartbeat is therefore the ESD times this scaling factor. The estimated chamber diastolic diameter is then calculated by dividing the estimated chamber systolic diameter by the fractional shortening. The stroke volume in picoliters is therefore:

SV=πchamberLengthchamberDiastolicRadius21000πchamberLengthchamberSystolicRadius21000

Cardiac output per second was calculated by integrating all stroke volumes that occurred in each second. The heart rate is the inverse of the duration between the initiation of adjacent contractions while contractions per second is the integral number of contractions the occurred in each second. The median heartbeat was derived by extracting the median diameter at each time point for time-aligned heartbeats until the median duration was reached. Quartiles were calculated similarly. Longer duration and more weakly relaxing heartbeats attenuate the apparent EDD at the end of the median heartbeat which is why a diameter mismatch between the initiation and end of the median heartbeat develops in animals exhibiting diastolic dysfunction. The heart wall velocity for each dataset was calculated using the derivative of the median heartbeat for that dataset. The time to peak velocity of contraction, and from peak contraction to peak relaxation, was calculated for each animal using their median heart wall velocities.

Heartbeat visualization

We generated all graphs using DataGraph 4β (Visual Data Tools) using the data output from Diadem 2011. Two-dimensional probability maps of chamber diameter and event duration were calculated in Diadem 2011 by aligning all contractions and heartbeats to the initiation of contraction and then displaying the normalized probability in DataGraph 4β. Probability maps of fractional shortening and systolic interval were similarly normalized to the maximum observed probability. Because sandman mutants displayed considerable phenotypic variability from animal to animal (see Figure 3A and Video 2), we did not find all datasets to exhibit normal distribution. Therefore, nearly all data is shown using Tukey boxplots so that the distributions can be accurately compared. The exception to this was when stroke volumes were juxtaposed with cardiac output per second; we presented stroke volumes as mean +/− s.d. for ease of visualization. Lastly, representative images of the heart, kymographs, and videos were overlaid with the heartbeat digitizations using ImageJ scripts.

Statistical analysis

To more accurately reflect sample size and variability, we considered individual heartbeats as interdependent and performed all statistics using the mean values for each animal. The letter n therefore denotes the number of independent biological experiments, detailed in Table 3. Pilot experiments established that the principal phenotypes and their rescue were sufficiently robust to be statistically tested using moderate sample sizes. Because not all datasets exhibited normal distributions, we utilized a non-parametric statistical test, Kruskal-Wallis followed by Dunn’s Multiple Comparison Test. ANOVA followed by Tukey’s multiple comparison test reported similar results without material deviations in significance. For the sensitivity tests to excitation light intensity and mounting duration presented in Figure 1—figure supplement 1, we utilized paired one-way ANOVA followed by Holm-Sidak’s multiple comparisons test. In all panels, significance is represented as ns = not significant, */#p<0.05, **/##p<0.01, ***/###p<0.001. Statistical tests were performed in Prism 6 software (GraphPad, La Jolla, CA). Fits were generated in DataGraph 4β using a linear least squares minimization method of the animal means, with outliers exceeding one and one-half the standard deviation excluded from the regression analysis.

Table 3

Intravital imaging sample sizes.

https://doi.org/10.7554/eLife.20851.024
Genotype and agenGenotype and agen
w1118 10 days10w1118 + R94C02 > sandman cDNA 50 days12
w1118 20 days12sandmanΔ + R94C02 > sandman cDNA 50 days27
w1118 30 days13sandmanΔ + tinCΔ4 > sandman cDNA 50 days12
w1118 50 days17sandmanΔ + R37B05 > sandman cDNA 50 days13
w1118 70 days12sandmanΔ / BSC267[Df] 50 days12
w1118 110 days13sandmanΔ / clean excision 50 days13
Canton S 10 days8sandmanΔ / BSC267[Df] 70 days15
Canton S 20 days11sandmanΔ / clean excision 70 days7
Canton S 30 days14tinCΔ4 control 40 days 25C15
Canton S 50 days13kk110628 / BSC431[Df] control 40 days 25C11
Canton S 70 days9tinCΔ4 > kk110628 / BSC431[Df] 40 days 25C11
Canton S 110 days14Canton S 30 days - vary light4
sandmanΔ 10 days13Canton S 40 days - mounting duration4
sandmanΔ 20 days9sandmanΔ 110 days + EGTA3
sandmanΔ 30 days14sandmanΔ 110 days + valinomycin3
sandmanΔ 50 days15wild-type 30 days - visualize sarcomeres9
sandmanΔ 70 days12sandmanΔ 30 days - visualize sarcomeres4

Electrophysiology

Patch pipettes with resistances of 3.5–5 MΩ were pulled from borosilicate glass capillaries (1.5 mm O.D, 0.86 mm I.D., Sutter Instruments, Novato, CA) using a P-1000 pipette puller (Sutter Instruments) and fire-polished using a microforge (MF-830, Narishige, Tokyo, Japan). Authenticated and mycoplama tested Chinese hamster ovary cells (CHO-K1), acquired from the University of California San Francisco Core Facility via the European Collection of Authenticated Cell Cultures (ECACC 85051005), were grown in F12-K media supplemented with 10% fetal bovine serum and passaged fewer than nine times. 70% confluent 30 mm petri dishes were transfected overnight with FuGene 6 (Promega, Madison, WI) using 1 µg total plasmid. The cells were then replated onto poly-L lysine coated coverslips and allowed to recover for two hours before recording. Pipettes were mounted onto a CV-7B headstage (Molecular Devices, Sunnyvale, CA) incorporating an Ag/AgCl electrode and attached to a MP-285 micromanipulator (Sutter Instruments). The data were lowpass filtered to 10 kHz using a Multiclamp 700 B amplifier and digitized at 50 kHz using a Digidata 1440 A analogue to digital convertor and pClamp 10 software (Molecular Devices). Analysis was performed off-line using Clampfit 10.5 (Molecular Devices) and visualized using DataGraph 4β (Visual Data Tools). The 200 ms voltage steps in whole cell mode were made in 20 mV increments ranging from −120 mV to +80 mV, from a holding potential of −80 mV. The internal pipet solution contained 150 K+, 5 Na+, 3 Mg2+, 161 Cl, 10 HEPES, pH 7.4 and the bath solution contained 5 K+, 150 Na+, 3 Mg2+, 1 Ca2+, 163 Cl-, 10 HEPES, pH 7.4 (in mM). Potassium selectivity experiments were also performed in the whole cell mode with an internal pipet solution containing 150 K+, 5 Na+, 3 Mg2+, 161 Cl-, 10 HEPES, pH 7.4 (in mM). The bath solution [K+] and [Na+] or [NMDG+] are as indicated in Figure 5B, with 3 Mg2+, 1 Ca2+, 161 Cl, 10 HEPES, pH 7.4, excepting the ‘Divalent-free’ solution which substitutes 2 EDTA for the divalent cations (in mM). The superfusion pipette had an internal diameter of 200 µm and the perfusate was gated using an Octaflow multi-valve perfusion system (ALA Scientific, Farmingdale, NY). Each solution was maintained for 10 sweeps of a voltage ramp protocol (200 ms, −120 mV to 80 mV ramp with a 300 ms 0 mV hold). This data was downsampled to 1 kHz and sweeps 2–10 were averaged for each condition, normalized to the peak current observed in the 5 K+ bath solution.

In vivo pharmacology

110-day-old male flies were imaged before and after bolus injection of 10 mM EGTA or 100 µM valinomycin (Sigma-Aldrich, St. Louis, MO) containing artificial hemolymph-like solution (AHLS): 113 Na+, 5 K+, 8.2 Mg2+, 2 Ca2+, 133 Cl, 5 HEPES, 4 HCO3, 1 H2PO4,, 10 Sucrose, 5 Trehalose, pH 7.5 (in mM). Borosilicate glass pipettes (1 mm OD, 0.75 mm ID, A-M Systems, Sequim, WA) were pulled using a P-1000 puller (Sutter Instruments). Pipette tip diameters of 50–75 µm were created by crushing the taper with forceps and visually confirming their diameter using a microforge (Narishige MF-830). Bolus injections into the abdomen of flies under ice anesthesia were approximately 1000 pL in volume and were made using a Femtojet (Eppendorf, Hamburg, Germany), with the pipette positioned using a manual micromanipulator (World Precision Instruments, Sarasota, FL).

Intravital imaging of sarcomere dynamics

Cardiomyocyte sarcomere dynamics in the posterior half of the second heart chamber of 30-day-old GFP α-actinin [CC01961] animals (Buszczak et al., 2007) were imaged using 19 mW per mm2 blue excitation light, delivered through a 40x/1.3 NA objective (Olympus UPLFLN40XO), with the excitation pattern again restricted to the field of view of the acquired image. Images were captured at 60 frames per second but with otherwise identical camera settings as above. The heart wall detection algorithm was adapted to trace single sarcomeres in YT kymographs of paired left and right cardiomyocytes. The data were subsequently analyzed by calculating the amplitude of coherence between the two sarcomeres:

Coherence=PositionLeftTimePositionRightTime

References

  1. 1
  2. 2
  3. 3
  4. 4
  5. 5
  6. 6
  7. 7
  8. 8
  9. 9
  10. 10
  11. 11
  12. 12
    Identification of heart rate-associated loci and their effects on cardiac conduction and rhythm disorders
    1. M den Hoed
    2. M Eijgelsheim
    3. T Esko
    4. BJ Brundel
    5. DS Peal
    6. DM Evans
    7. IM Nolte
    8. AV Segrè
    9. H Holm
    10. RE Handsaker
    11. HJ Westra
    12. T Johnson
    13. A Isaacs
    14. J Yang
    15. A Lundby
    16. JH Zhao
    17. YJ Kim
    18. MJ Go
    19. P Almgren
    20. M Bochud
    21. G Boucher
    22. MC Cornelis
    23. D Gudbjartsson
    24. D Hadley
    25. P van der Harst
    26. C Hayward
    27. M den Heijer
    28. W Igl
    29. AU Jackson
    30. Z Kutalik
    31. J Luan
    32. JP Kemp
    33. K Kristiansson
    34. C Ladenvall
    35. M Lorentzon
    36. ME Montasser
    37. OT Njajou
    38. PF O'Reilly
    39. S Padmanabhan
    40. B St Pourcain
    41. T Rankinen
    42. P Salo
    43. T Tanaka
    44. NJ Timpson
    45. V Vitart
    46. L Waite
    47. W Wheeler
    48. W Zhang
    49. HH Draisma
    50. MF Feitosa
    51. KF Kerr
    52. PA Lind
    53. E Mihailov
    54. NC Onland-Moret
    55. C Song
    56. MN Weedon
    57. W Xie
    58. L Yengo
    59. D Absher
    60. CM Albert
    61. A Alonso
    62. DE Arking
    63. PI de Bakker
    64. B Balkau
    65. C Barlassina
    66. P Benaglio
    67. JC Bis
    68. N Bouatia-Naji
    69. S Brage
    70. SJ Chanock
    71. PS Chines
    72. M Chung
    73. D Darbar
    74. C Dina
    75. M Dörr
    76. P Elliott
    77. SB Felix
    78. K Fischer
    79. C Fuchsberger
    80. EJ de Geus
    81. P Goyette
    82. V Gudnason
    83. TB Harris
    84. AL Hartikainen
    85. AS Havulinna
    86. SR Heckbert
    87. AA Hicks
    88. A Hofman
    89. S Holewijn
    90. F Hoogstra-Berends
    91. JJ Hottenga
    92. MK Jensen
    93. A Johansson
    94. J Junttila
    95. S Kääb
    96. B Kanon
    97. S Ketkar
    98. KT Khaw
    99. JW Knowles
    100. AS Kooner
    101. JA Kors
    102. M Kumari
    103. L Milani
    104. P Laiho
    105. EG Lakatta
    106. C Langenberg
    107. M Leusink
    108. Y Liu
    109. RN Luben
    110. KL Lunetta
    111. SN Lynch
    112. MR Markus
    113. P Marques-Vidal
    114. I Mateo Leach
    115. WL McArdle
    116. SA McCarroll
    117. SE Medland
    118. KA Miller
    119. GW Montgomery
    120. AC Morrison
    121. M Müller-Nurasyid
    122. P Navarro
    123. M Nelis
    124. JR O'Connell
    125. CJ O'Donnell
    126. KK Ong
    127. AB Newman
    128. A Peters
    129. O Polasek
    130. A Pouta
    131. PP Pramstaller
    132. BM Psaty
    133. DC Rao
    134. SM Ring
    135. EJ Rossin
    136. D Rudan
    137. S Sanna
    138. RA Scott
    139. JS Sehmi
    140. S Sharp
    141. JT Shin
    142. AB Singleton
    143. AV Smith
    144. N Soranzo
    145. TD Spector
    146. C Stewart
    147. HM Stringham
    148. KV Tarasov
    149. AG Uitterlinden
    150. L Vandenput
    151. SJ Hwang
    152. JB Whitfield
    153. C Wijmenga
    154. SH Wild
    155. G Willemsen
    156. JF Wilson
    157. JC Witteman
    158. A Wong
    159. Q Wong
    160. Y Jamshidi
    161. P Zitting
    162. JM Boer
    163. DI Boomsma
    164. IB Borecki
    165. CM van Duijn
    166. U Ekelund
    167. NG Forouhi
    168. P Froguel
    169. A Hingorani
    170. E Ingelsson
    171. M Kivimaki
    172. RA Kronmal
    173. D Kuh
    174. L Lind
    175. NG Martin
    176. BA Oostra
    177. NL Pedersen
    178. T Quertermous
    179. JI Rotter
    180. YT van der Schouw
    181. WM Verschuren
    182. M Walker
    183. D Albanes
    184. DO Arnar
    185. TL Assimes
    186. S Bandinelli
    187. M Boehnke
    188. RA de Boer
    189. C Bouchard
    190. WL Caulfield
    191. JC Chambers
    192. G Curhan
    193. D Cusi
    194. J Eriksson
    195. L Ferrucci
    196. WH van Gilst
    197. N Glorioso
    198. J de Graaf
    199. L Groop
    200. U Gyllensten
    201. WC Hsueh
    202. FB Hu
    203. HV Huikuri
    204. DJ Hunter
    205. C Iribarren
    206. B Isomaa
    207. MR Jarvelin
    208. A Jula
    209. M Kähönen
    210. LA Kiemeney
    211. MM van der Klauw
    212. JS Kooner
    213. P Kraft
    214. L Iacoviello
    215. T Lehtimäki
    216. ML Lokki
    217. BD Mitchell
    218. G Navis
    219. MS Nieminen
    220. C Ohlsson
    221. NR Poulter
    222. L Qi
    223. OT Raitakari
    224. EB Rimm
    225. JD Rioux
    226. F Rizzi
    227. I Rudan
    228. V Salomaa
    229. PS Sever
    230. DC Shields
    231. AR Shuldiner
    232. J Sinisalo
    233. AV Stanton
    234. RP Stolk
    235. DP Strachan
    236. JC Tardif
    237. U Thorsteinsdottir
    238. J Tuomilehto
    239. DJ van Veldhuisen
    240. J Virtamo
    241. J Viikari
    242. P Vollenweider
    243. G Waeber
    244. E Widen
    245. YS Cho
    246. JV Olsen
    247. PM Visscher
    248. C Willer
    249. L Franke
    250. J Erdmann
    251. JR Thompson
    252. A Pfeufer
    253. N Sotoodehnia
    254. C Newton-Cheh
    255. PT Ellinor
    256. BH Stricker
    257. A Metspalu
    258. M Perola
    259. JS Beckmann
    260. GD Smith
    261. K Stefansson
    262. NJ Wareham
    263. PB Munroe
    264. OC Sibon
    265. DJ Milan
    266. H Snieder
    267. NJ Samani
    268. RJ Loos
    269. Global BPgen Consortium
    270. CARDIoGRAM Consortium
    271. PR GWAS Consortium
    272. QRS GWAS Consortium
    273. QT-IGC Consortium
    274. CHARGE-AF Consortium
    (2013)
    Nature Genetics 45:621–631.
    https://doi.org/10.1038/ng.2610
  13. 13
  14. 14
  15. 15
  16. 16
  17. 17
  18. 18
  19. 19
  20. 20
  21. 21
  22. 22
  23. 23
  24. 24
  25. 25
  26. 26
  27. 27
  28. 28
  29. 29
  30. 30
  31. 31
  32. 32
  33. 33
  34. 34
  35. 35
  36. 36
  37. 37
  38. 38
  39. 39
  40. 40
  41. 41
  42. 42
  43. 43
  44. 44
  45. 45
  46. 46
  47. 47
  48. 48
  49. 49
  50. 50
  51. 51
  52. 52
  53. 53
  54. 54
  55. 55
  56. 56
  57. 57
  58. 58
  59. 59
  60. 60
  61. 61
  62. 62

Decision letter

  1. Talila Volk
    Reviewing Editor; Weizmann Institute of Science, Israel

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Age-dependent diastolic heart failure in an in vivo Drosophila model" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Naama Barkai as the Senior Editor. The following individual involved in review of your submission has agreed to reveal his identity: Chao Zhou (Reviewer #3).

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

Summary:

Klassen et al., describe the development of a novel method to analyze dynamic heart contraction in live adult fly based on high resolution fluorescent imaging. Using this method, they show that, like in humans, the aging Drosophila heart manifests primarily defects in relaxation (diastole) rather than in heart contraction (systole). Using their novel method the authors discovered that a pair of two-pore potassium channel (K2P) subunits, largely dispensable in younger flies, are necessary for terminating contraction (systole) in aged animals, where their loss culminates in fibrillatory cardiac arrest. This study paves the road for using the fly heart as a model for understanding the signaling networks maintaining cardiac performance during normal aging.

Essential revisions:

1) Authors need to show that sandman and galene are expressed in the adult fly heart.

2) Does overexpression of sandman "rescue" the aging phenotype?

3) Need a negative control for galenesandman interaction in cell culture. Since other K2P channels were screened and had no effects on heart function they should be tested.

Figure 1 – Because it has been shown in previous studies of the fly heart that there are differences in the rates and rhythmicity of anterograde and retrograde hearts beats in intact flies one would expect that heartbeat parameters would also vary. Was the data shown in Figure 1F, G based on all the heartbeats in a record or subsets of the data binned into anterograde or retrograde? This is not clear nor are the differences in the two types of contractions discussed with respect to the data.

Subsection “Imaging cardiac performance in intact, unanesthetized Drosophila”, last paragraph – The authors state that irradiation has no effect on heart function at low levels, however, they cannot actually show what the heart is doing without UV irradiation. In our experience UV irradiation increases heart rate and arrhythmicity. How would they address this?

Subsection “Imaging cardiac performance in intact, unanesthetized Drosophila”, last paragraph – Since it has been reported that 80% of female flies are dead after 18 hours of desiccation, it is not clear how the preparation can still be viable after 19 hours glued to a slide.

Subsection “Normal aging manifests as diastolic dysfunction while preserving contractile performance”, second paragraph – The authors group data from the two 'wild-type' genetic backgrounds, w1118 and Canton S, for analysis. This is troublesome as these two genotypes have very different genetic backgrounds – known to influence heart function. The average fly size and heart size in these two genotypes are different. Having similar rates does not seem sufficient rational for grouping data from these two lines. Especially since they are also using this data to generate other parameters that are not based solely on rate (e.g. Cardiac output).

Subsection “A pair of K2P subunits, sandman and galene, are required for terminating systole in aged animals” – Why were two different cardiac drivers used for the sandman and galene KD? What happens when the two drivers are reversed, does tinCd4>Sandman RNAi have the same effect as for R94C02>Sandman KD? And vice versa for Galene RNAi?

Subsection “Sandman is required to maintain diverse biological oscillators”, first paragraph – "oscillatory balance degenerates catastrophically with age" What does this mean in terms of cardiomyocyte function how does it relate to the persistent contractions in sandman KD?

Introduction, third paragraph – The authors misrepresent their system as a high throughput system. If it is high throughput why did they have to pool wt genotypes and analyze only 24 flies per age group for the aging study? In fact, as for all other fly heart model systems including OCT, their system requires manual mounting and orientation of flies and manual tracing of data after the fact to facilitate analysis. These are not characteristics of a high throughput system.

4) The work has one major limitation, however. It is inferred that calcium cycling abnormalities underlie the actual relaxation abnormalities detected, but cardiomyocyte calcium cycling is not measured. This seems curious as the proposed role of calcium is central to the overall proposed mechanism introduced in the last paragraph of the subsection “The sandman phenotype reflects a loss of repolarization rather than structurally congestive remodeling”. Moreover, others have successfully measured cardiomyocyte calcium cycling in intact Drosophila using fluorescent techniques that should be readily adaptable to the current system (for example, the Maack group's work on Drosophila mitochondrial-SR calcium microdomains in Circ Res 2012). Direct calcium measurements are not replaceable by inferences drawn after introduction of EGTA, etc., and their absence here substantially limits confidence in the overall mechanistic conclusions.

https://doi.org/10.7554/eLife.20851.028

Author response

Essential revisions:

1) Authors need to show that sandman and galene are expressed in the adult fly heart.

We have microdissected the adult fly heart from the rest of the animal and utilized RT-PCR to confirm that sandman and galene transcripts are indeed expressed there as expected (Figure 3—figure supplement 1).

2) Does overexpression of sandman "rescue" the aging phenotype?

We have overexpressed the sandman cDNA in the heart of wild type and sandman mutant animals through 50 days of age, when wild type animals show signs of cardiac decline and sandman mutants show profound defects in heart function. Overexpression robustly rescues the sandman mutant phenotype (Figures 3, 4, and Figure 4—figure supplement 2) but does not appear to improve or degrade cardiac performance in wild-type animals (Figures 3, and Figure 4—figure supplement 2).

3) Need a negative control for galene – sandman interaction in cell culture. Since other K2P channels were screened and had no effects on heart function they should be tested.

As a negative control for heteromeric interaction in cell culture, we recorded from CHO cells combinatorially expressing the K2P genes most closely related to sandman and galene, CG1688 and CG10864, to determine whether these subunits might form functional homomers or, if not, whether they were capable of forming functional heteromeric channels if co-expressed with sandman or galene. We found that, of all possible combinations of these genes, only sandman-galene co-transfection produced measurable currents (Figure 5 and Figure 5—figure supplement 1).

Figure 1 – Because it has been shown in previous studies of the fly heart that there are differences in the rates and rhythmicity of anterograde and retrograde hearts beats in intact flies one would expect that heartbeat parameters would also vary. Was the data shown in Figure 1F, G based on all the heartbeats in a record or subsets of the data binned into anterograde or retrograde? This is not clear nor are the differences in the two types of contractions discussed with respect to the data.

All data combine anterograde and retrograde heartbeats. We do observe qualitative differences in the contraction-relaxation waveform of anterograde versus retrograde heartbeats (e.g. Figure 1E), which may reflect differences in pressure waves arising from bolus flow and/or differences in the electrical waveform emanating from the two distinct pacemakers. Although we do plan to extend our computer code to automatically discriminate anterograde and retrograde heartbeats in the future, we do not believe that subdividing our dataset is necessary to support the principal conclusions of our present manuscript. Sustained contractions appear for both directions in sandman mutants that are not yet terminally contracted (e.g. Video 2). Furthermore, contraction-relaxation waves become discontinuous and chaotic in sandman mutants or galene knockdown, which makes any segmentation of anterograde and retrograde heartbeats less trivial than two-point discrimination. Because of these observations, we feel that attempting to subdivide the data into two separate bins would not increase our mechanistic insight and risks diminishing the readability of the manuscript and particularly the figures.

Subsection “Imaging cardiac performance in intact, unanesthetized Drosophila”, last paragraph – The authors state that irradiation has no effect on heart function at low levels, however, they cannot actually show what the heart is doing without UV irradiation. In our experience UV irradiation increases heart rate and arrhythmicity. How would they address this?

At the light intensities utilized throughout the paper, we did not observe any meaningful arrhythmicity in wild-type animals. It is possible that spatially restricting illumination to a small region of interest and not using UV excitation has avoided the potential arrhythmic effects previously observed by the reviewer.

We do not use UV irradiation during imaging. We only utilize a brief 10s pulse of 405nm LED light to polymerize the optical glue during cold anesthesia. After 15 minutes of recovery, imaging is performed with green light excitation (538-568nm, 4.3 mW/mm2) limited to a small (150x500um) region of interest, which is less than 20% of the total surface of the abdomen. Fly hearts imaged using optical coherence tomography (e.g. Wolf et al., 2006; Alex et al., 2015) or near-IR spectroscopy (Wasserthal, 2007), neither of which utilize UV-activated optical cement, exhibit similar mean heart rates to those reported here (4.5-6.5 Hz vs 5.6 Hz in 10 day animals declining to 3.8 Hz at 110 days).

We tested the effects of increasing green light excitation on the heart and found a 0.39% increase in heart rate per mW/mm2 (R2 = 0.93) which would suggest that our green light illumination levels increase the heart rate less than 2% (Figure 1—figure supplement 1A). To ensure that we minimized any effects arising from the excitation light, we also tested illumination intensities 60% lower than utilized, equivalent to a small spot of restricted light only 50% stronger than sunlight, or less total irradiation than the fly would experience exposed during midday. We observed no differences between these two intensities, suggesting that we minimized any excitation light related artifacts. Light intensities 20-fold higher than utilized exhibited significant tachycardia but only the occasional early aftercontraction.

Subsection “Imaging cardiac performance in intact, unanesthetized Drosophila”, last paragraph – Since it has been reported that 80% of female flies are dead after 18 hours of desiccation, it is not clear how the preparation can still be viable after 19 hours glued to a slide.

We hydrated the immobilized flies overnight using a small drinking capillary and apologize for omitting this detail, now corrected.

Subsection “Normal aging manifests as diastolic dysfunction while preserving contractile performance”, second paragraph – The authors group data from the two 'wild-type' genetic backgrounds, w1118 and Canton S, for analysis. This is troublesome as these two genotypes have very different genetic backgrounds – known to influence heart function. The average fly size and heart size in these two genotypes are different. Having similar rates does not seem sufficient rational for grouping data from these two lines. Especially since they are also using this data to generate other parameters that are not based solely on rate (e.g. Cardiac output).

Our conclusion that systolic function is preserved while the kinetics of relaxation are selectively affected during aging is true for both Canton S and w1118 genotypes (see Author response image 1). The data were combined because, in our experiments, we found them comparable.

The Canton S background is the preferred genetic background for behavioral studies, while white and particularly yellow mutants are known to display behavioral defects in certain assays. We retained the w1118 allele to facilitate building and confirming transgenic backgrounds but outcrossed it for six generations to our Canton S stock to eliminate any unlinked polymorphisms. We collected the Canton S and w1118 datasets to confirm that the retention of w1118 did not grossly affect cardiac rhythmicity or output. This combined dataset was only used for Figure 2; all other figures utilized the w1118 dataset alone as all subsequent genetics were performed in that background.

To clarify, all heart parameters are calculated on a heartbeat by heartbeat basis for each fly and do not rely on estimates or parameters generated at the population level in any way. Therefore, if there were material differences in heart function in Canton S vs. w1118 animals, it would manifest in the Tukey boxplots.

If one scrutinizes these datasets separately, they do reveal that the w1118 dataset has a modest trend towards higher cardiac output (+8.5%) resulting from higher stroke volume, partially mitigated by lower heart rate (see Figure above). The genesis of this difference is unclear but may reflect the fact that our Canton S animals are somewhat healthier and are therefore easier to subtly overcrowd during rearing, leading to slightly smaller animals.

It is possible that polymorphisms not linked to the white locus have contributed to the differences observed by others, but their genetic identity or prevalence within stocks across the fly community has not been characterized in the literature.

Subsection “A pair of K2P subunits, sandman and galene, are required for terminating systole in aged animals” – Why were two different cardiac drivers used for the sandman and galene KD? What happens when the two drivers are reversed, does tinCd4>Sandman RNAi have the same effect as for R94C02>Sandman KD? And vice versa for Galene RNAi?

We utilized tinCd4::Gal4 for all RNAi experiments. It is the strongest heart Gal4 driver and is almost universally used by our peers for heart-specific RNAi (e.g. Neely et al., 2010). We initially used R94C02 for rescue out of genetic convenience given the stocks we had assembled at the time. We have now attempted rescuing the sandman phenotype using two additional Gal4s to allay the concerns expressed by the reviewer. tinCd4::Gal4 robustly rescues the sandman phenotype while a Gal4 expressed in many muscle subtypes excepting the heart (R37B05::Gal4) does not rescue (Figure 4—figure supplement 3).

Subsection “Sandman is required to maintain diverse biological oscillators”, first paragraph – "oscillatory balance degenerates catastrophically with age" What does this mean in terms of cardiomyocyte function how does it relate to the persistent contractions in sandman KD?

The term “oscillatory balance” was meant to reflect the balance between contraction and relaxation that progressively trends toward persistent contractions in aged sandman mutants. This is most easily observed in Figure 4—figure supplement 1B, where the median heart contraction is not balanced with an equivalent magnitude heart relaxation in aged sandman mutants. We have clarified this language in the manuscript.

Introduction, third paragraph – The authors misrepresent their system as a high throughput system. If it is high throughput why did they have to pool wt genotypes and analyze only 24 flies per age group for the aging study? In fact, as for all other fly heart model systems including OCT, their system requires manual mounting and orientation of flies and manual tracing of data after the fact to facilitate analysis. These are not characteristics of a high throughput system.

Our apologies. In our defense, we routinely mount and record 15 flies, collecting approximately 7,000 heartbeats per hour, and our heart wall detection and heartbeat segmentation algorithms are fully automated. We believe this is a significant throughput advance over existing methodologies for any heart system. The only manual steps are selecting the longitudinal position for the kymograph and tracing the chamber once to estimate its average systolic volume, which is script assisted and takes less than 30 seconds per fly.

The wild-type dataset comprises 146 flies in total. We combined the w1118 and Canton S datasets because they were not meaningfully different from one another and were available for inclusion. We did not collect additional flies for this dataset as the principal conclusions were readily quantifiable at this sample size. We strongly believe that our methodology scales particularly well and will permit considerable forward genetic screening in future studies but have moderated references to throughput in line with the reviewer’s comments.

4) The work has one major limitation, however. It is inferred that calcium cycling abnormalities underlie the actual relaxation abnormalities detected, but cardiomyocyte calcium cycling is not measured. This seems curious as the proposed role of calcium is central to the overall proposed mechanism introduced in the last paragraph of the subsection “The sandman phenotype reflects a loss of repolarization rather than structurally congestive remodeling”. Moreover, others have successfully measured cardiomyocyte calcium cycling in intact Drosophila using fluorescent techniques that should be readily adaptable to the current system (for example, the Maack group's work on Drosophila mitochondrial-SR calcium microdomains in Circ Res 2012). Direct calcium measurements are not replaceable by inferences drawn after introduction of EGTA, etc., and their absence here substantially limits confidence in the overall mechanistic conclusions.

Unfortunately, there are no published methodologies for measuring cardiomyocyte calcium cycling in intact Drosophila. We agree that it would be useful to monitor the calcium and voltage waveforms in vivo but the field has yet to surmount technical limitations associated with these experiments. The calcium wave observed by the Maack group and others utilizes the in situ preparation which beats at approximately 1-2Hz and is therefore sufficiently slow for GCaMP calcium imaging. However, this calcium wave is several fold longer in duration than what one would expect to observe in vivo, which may arise from the heart being removed from its normal physiochemical environment and/or slow GCAMP off kinetics distorting the signal. The fly heart beats at approximately 5-6 Hz in vivo, which means that best current generation genetically encoded calcium indicators do not have sufficiently rapid off-decay times (t1/2 GCaMP6f 350ms, Chen et al., Nature 2013) to adequately resolve individual calcium waves in vivo. Given such dramatic differences in the calcium waveform in situ vs. in vivo, it stands to reason that there exist significant differences in the relative strength and kinetics of the various inward and outward conductances, making comparisons about ion channels problematic.

We invested considerable time attempting to overcome this technical hurdle, even testing GCAMP6 variants with reduced calcium affinity, but have determined that the temporal resolution of the optical signals is currently inadequate, compounded by the difficulty imaging cytosolic GFP signals intravitally and addressing motion artifacts that cannot be deconvolved without ratiometric approaches. This last point of particular concern as most published papers using this methodology neglect to compensate for motion artifacts arising from the contraction-relaxation cycle, which Lin et al., 2011, who developed this methodology for the Drosophila heart, demonstrated represents 50% of the resultant signal (see their Supplementary Figure 1). Effectively, one can derive an equivalent ‘calcium’ signal using cytosolic GFP. Cetoxymethyl ester conjugated calcium sensitive dyes, while having sufficiently fast off-kinetics and internal ratiometric controls, have been reported by others to load very poorly in Drosophila (e.g. Lin et al., 2011 and our experience). We tried paralyzing the heart in situ using blebbistatin and/or cytochalasin-D but have found that these compounds trigger significant dysrhythmia in wild-type animals, as measured by genetically-encoded voltage indicators.

While we are sympathetic to the reviewer’s desire to directly visualize calcium, we feel that the experiments detailed in Figure 6 should not be discounted as inadequately supporting our model as it relates to the scope of this publication. We are unaware of any mechanistic basis for a calcium independent contraction-relaxation of the sarcomere but have moderated our text accordingly.

To better understand the terminal phenotype of sandman mutants, we directly visualized sarcomere dynamics in vivo, for which the biophysical linkage between calcium and contraction is well established. This was a significant technical accomplishment and provides excellent spatiotemporal resolution of individual sarcomeric behavior (Video 3). During the sustained contractions observed in sandman mutants, sarcomeres actively contract and relax out of phase with one another, suggesting that sufficient ATP was available locally to drive myosin activity, myosin dissociation and the various pumps that can sequester calcium. Interestingly, this fibrillatory behavior periodically and rapidly resolves itself in a synchronous manner globally (Figure 6, Video 3), an effect that is most plausibly explained by a global repolarization closing sarcolemmal Cav channels, which inactivate much more slowly in Drosophila relative to vertebrates (t1/2 1-2s, Haraet al., J Neurogenetics 2015) and could therefore generate a prolonged window current sustaining calcium influx in the absence of sufficient repolarization back to the resting potential.

Further support of our model comes from our pharmacological experiments, which importantly are acute (seconds). First, chelating extracellular calcium using membrane impermeable EGTA demonstrates that calcium influx across the sarcolemmal membrane is necessary for sustaining contractions in sandman mutant animals. Second, creating an artificial but highly selective K+ leak across the sarcolemmal membrane using valinomycin also terminated the persistent contractions. Accordingly, these experiments exclude a calcium and/or potassium independent means of sustaining the persistent contractions in sandman mutants. Because potassium channels play well-established roles in cardiac repolarization back to the resting potential and L-type voltage gated Ca channels are essential for heart contractions in Drosophila, we feel that a mechanistic link between the loss of a potassium channel and the persistent contractions observed is not unduly speculative.

What we do not yet understand is why the phenotype is progressively age-dependent and what mechanisms cause the fibrillatory cycling of individual sarcomeres during the sustained contractions. Having high-resolution voltage indicators as well as calcium measures for the cytosol and SR would allow us to address these questions, which we hope to develop in a future manuscript. In conclusion, we feel that our intravital imaging methodology and our characterization of this novel potassium channel are worthy of publication and look forward to extending this work in future studies.

https://doi.org/10.7554/eLife.20851.029

Article and author information

Author details

  1. Matthew P Klassen

    Department of Physiology, Howard Hughes Medical Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    MPK, Conceptualization, Software, Formal analysis, Funding acquisition, Investigation, Visualization, Methodology, Writing—original draft
    For correspondence
    matthew.klassen@ucsf.edu
    Competing interests
    The authors declare that no competing interests exist.
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3633-2512
  2. Christian J Peters

    Department of Physiology, Howard Hughes Medical Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    CJP, Conceptualization, Funding acquisition, Investigation, Methodology, Writing—review and editing
    Competing interests
    The authors declare that no competing interests exist.
  3. Shiwei Zhou

    Department of Physiology, Howard Hughes Medical Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    SZ, Investigation, Methodology, Writing—review and editing
    Competing interests
    The authors declare that no competing interests exist.
  4. Hannah H Williams

    Department of Physiology, Howard Hughes Medical Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    HHW, Investigation, Methodology, Writing—review and editing
    Competing interests
    The authors declare that no competing interests exist.
  5. Lily Yeh Jan

    1. Department of Physiology, Howard Hughes Medical Institute, University of California, San Francisco, San Francisco, United States
    2. Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, United States
    Contribution
    LYJ, Conceptualization, Supervision, Funding acquisition, Writing—review and editing
    Competing interests
    The authors declare that no competing interests exist.
  6. Yuh Nung Jan

    Department of Physiology, Howard Hughes Medical Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    YNJ, Conceptualization, Supervision, Funding acquisition, Writing—review and editing
    For correspondence
    YuhNung.Jan@ucsf.edu
    Competing interests
    The authors declare that no competing interests exist.

Funding

Jane Coffin Childs Memorial Fund for Medical Research (Postdoctoral Fellowship)

  • Matthew P Klassen

Heart and Stroke Foundation of Canada (Junior Personnel Fellowship)

  • Christian J Peters

National Institute of Mental Health (R37MH0653354)

  • Lily Yeh Jan

Howard Hughes Medical Institute (Investigators)

  • Lily Yeh Jan
  • Yuh Nung Jan

National Institutes of Health (R37NS040929)

  • Yuh Nung Jan

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

Acknowledgements

We thank M Petkovic, S Younger, S Barbel and T Cheng for technical support, W Zhang, M Petkovic and S Headland for critical reading of the manuscript, and members of the Jan laboratory for discussion. This work was supported by National Institutes of Health grant (R37NS040929) to YNJ and National Institutes of Mental Health grant (R37MH0653354) to LYJ. MK was supported by a Jane Coffin Childs fellowship and CJP by a junior personnel fellowship from the Heart and Stroke Foundation of Canada. We would also like to acknowledge the Drosophila Genomics Resource Center, supported by NIH grant 2P40OD010949-10A1. YNJ and LYJ are investigators of the Howard Hughes Medical Institute.

Reviewing Editor

  1. Talila Volk, Weizmann Institute of Science, Israel

Publication history

  1. Received: August 26, 2016
  2. Accepted: February 11, 2017
  3. Version of Record published: March 22, 2017 (version 1)

Copyright

© 2017, Klassen et al.

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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