TcCAKC knockouts were obtained by sequential allelic replacement by homologous recombination. Recombination cassettes were obtained by PCR of neomycin or hygromycin resistance genes flanked by 500 bp of TcCAKC 5′ and 3′ UTRs. The fragments were amplified with allele specific primers and cloned into TOPO-Blunt II vector. Constructs verified by sequencing where purified, linearized, and transfected into CL strain epimastigotes using AMAXA nucleofector system protocol U-033. Selection was carried out with 250 μg/ml of G418 and 100 μg/ml of hygromycin. Once selection was complete, parasite populations were subcloned by serial dilution and screened by PCR to verify the correct insertion of the replacement cassettes using primers annealing upstream and downstream of TcCAKC UTRs.

Level of expression of TcCAKC in single-allele replacement (sKO) and double-allele replacement parasites (dKO) was evaluated by qPCR. Epimastigotes were collected during mid-log phase of growth, washed once with 1x PBS pH 7.4 and homogenized in TRI Reagent^®. Total mRNA was extracted following the manufacturer protocol (Sigma-Aldrich, St. Louis, MO) followed by chloroform/ ethanol precipitation cDNA was obtained using SuperScript^® III First-Strand Synthesis System (ThermoFisher Scientific, Inc., Waltham, MA) and oligo-dT₍₂₀₎ primers. cDNA was analyzed by qPCR with Power SYBR Green PCR Master Mix (Applied Biosystems) and primers forward 5′ GAACGTGGTCGGGTCAATCT 3′ and reverse 5′ GAGGCGACGTGTGTGAGAAT 3′. All qPCR results were normalized against GAPDH and tubulin as housekeeping gene and indicated as ΔΔCq Mean ± SD of at least 3 independent experiments in triplicate.

CL strain epimastigotes were cultured in LIT media supplemented with 10% inactivated Fetal bovine serum (FBS) at 28°C. Knockout parasites were maintained with 250 μg/mL G418 (sKO) plus 100 μg/mL hygromycin (dKO) (Bone and Steinert, 1956). To evaluate the growth of the parasites, cells were diluted to a concentration of 1 × 10⁶/mL in LIT media supplemented with FBS and antibiotics and counted every 24 h for 5 days in a Z2 Cell Counter (Beckman Instruments). Cell counts were taken in triplicate from three independent experiments. Statistics were ran using one factor ANOVA with post-hoc Bonferroni test to compare mutant and wild-type (WT) cells. To evaluate the infective capacity of the mutants, differentiation to metacyclic trypomastigote forms was induced under chemically defined conditions using triatomine artificial urine (TAU) medium as described (Contreras et al., 1985). Epimastigotes at 4 days of growth were collected by centrifugation at 1,600 × g for 10 min, washed once in phosphate buffered saline solution (PBS) pH 7.4, resuspended in TAU media and incubated 2 h at 28°C. The supernatant was collected and resuspended in TAU with amino acids (TAU3AAG), incubated for up to 7 days at 28°C, collected by centrifugation resuspended in 5 mL of Dulbecco's Modified Eagles Media (DMEM) supplemented with 20% fresh FBS to eliminate residual epimastigotes.

HEK-293 cells were plated onto coverlips in 12 well plates (1,000 cells/well) and incubated in supplemented HG-DMEM overnight at 37°C with 5% CO₂. Infections were performed at a multiplicity of infection (MOI) of 25:1 with either WT, sKO or dKO mutant trypomastigotes. After 6 h, the cells were washed 3 times with Hank's media and fresh DMEM was added. Coverslips were fixed in 4% paraformaldehyde-PBS at 6, 24, and 48 h, stained with DAPI (5 μg/ml) and mounted in Fluoromont media for quantification of intracellular parasites. All infection quantifications were done in 4 coverslips per experiment, in 3 or more independent experiments. At least 100 host cells were quantified per coverslip. The number of host cells vs. parasites was compared by Student t-test.

Aliquots of 1 × 10⁸ cells were diluted in Standard buffer (in mM: NaCl 135, KCl 5, CaCl₂ 1, MgSO₄ 1, glucose 5, HEPES 10 pH 7.4) plus 1 μM DisBac₂(3). Fluorescence was recorded at 1 Hz with excitation at 530 nm and emission at 560 nm. Calibration was performed by adding 1 μM of gramicidin to WT epimastigotes in NMDG Buffer (composition) and increasing concentrations of K⁺ gluconate (0.1, 1, 2, 5, 10, 25, 50, 100 mM). Calibration potentials were calculated using the theoretical Nernst potential equation [Veq=RTzFln([K+]out[K+]in)], where [K⁺]_in was assumed to be 120 mM (Van Der Heyden and Docampo, 2002). A linear line of best fit was generated and the equation for this line was used for interpolation of experimental data. Experimental measurements were done as described above. Resting membrane potential was averaged over the first 100 s of baseline recording. All recordings were done in Standard buffer unless otherwise indicated.

Epimastigotes were loaded with 6 μM BCECF-AM at 30°C for 30 min in standard buffer, washed twice and resuspended at a concentration of 1 × 10⁹ cells/mL in Standard buffer. Fluorescence was measured with excitation wavelengths of 490/440 nm and emission of 530 nm. Recordings were done in Standard Buffer unless otherwise indicated. Calibration was done in high K⁺ Standard Buffer (135 mM KCl, 1 mM MgSO₄, 1 mM CaCl₂, 5 mM glucose and 10 mM HEPES-Tris, pH 7.4), with 1 μM nigericin at various pHs (6, 6.5, 7.0, 7.4, 7.6, 8.0). Once stabilized, the fluorescent reading for any pH was averaged over 100 s and plotted against pH. The linear fit was then used to interpolate the experimental data.

Measurements of proton extrusion were done in the presence of 0.38 μM BCECF Free acid mixed with 1 × 10⁸ epimastigotes in low buffer standard solution (in mM: NaCl 135, KCl 5, CaCl₂ 1, MgSO₄ 1, glucose 5, HEPES 0.1 pH 7.4) (Benchimol et al., 1998). The recordings were done at excitation wavelengths of 490/440 nm and emission of 530 nm. Calibration was performed as indicated above. Differences in proton extrusion were measured by looking at differences in the slope over the first 50 s of recording (Initial rate of extrusion) or in the last 200 s of recording (Final). All experiments were done in low buffered standard buffer unless otherwise indicated.

Epimastigotes were loaded with 5 μM Fura2-AM (Molecular Probes) in BAG for 30 min at 30°C, washed twice and resuspended in BAG at a concentration of 5 × 10⁸ cells/ml. Aliquots of 5 × 10⁷ cells were taken for each measurement with excitation at 340/380 nm and emission at 525 nm. Recordings were performed in BAG unless otherwise indicated. Calibration was done by permeabilizing cells in BAG + 1 mM EGTA and then adding increasing concentrations of CaCl₂. The concentration of free calcium available was calculated using MaxChelator software (https://somapp.ucdmc.ucdavis.edu/pharmacology/bers/maxchelator/CaEGTA-TS.htm) and the Kd was calculated according to the manufacturer protocol. Experimental recordings were allowed to stabilize at baseline before addition of 1.8 mM CaCl₂ and averaged for 100 s after stabilization. For all the experiments, ionic replacement was done by substituting Na⁺, K⁺ or both by N-Methyl-D-glucamine (NMDG) in the corresponding standard buffer.

To demonstrate the function of TcCAKC as a potassium channel we expressed the protein in S. cerevisiae PLY246 (trk1Δ trk2Δ and tok1Δ null mutant). In this strain, the principal K⁺ permeation pathways have been eliminated and the cells require the supplementation of the media with high amounts of this ion to sustain their growth (Bertl et al., 2003). After 2 h of induction TcCAKC was expressed in the yeast vacuole and at later points (24 h) in the plasma membrane (Figure S3A). Importantly, the channel expression was able to revert the growth phenotype of this mutant providing evidence that it is, in fact, a K⁺ permeable channel (Figure S3B).

Two-electrode voltage clamp recodings of X. laevis oocytes expressing TcCAKC showed a stable resting membrane potential of −22.8 ± 7 mV (n = 24), similar to the membrane potential of control oocytes injected with DEPC water (−26.8 ± 6 mV, n = 24), indicating that the expression of the channel did not significantly affect the health of the oocytes. When cells where subjected to a voltage step protocol from −80 to 40 mV with a holding potential of −60 mV, oocytes expressing TcCAKC (Figure 2A black line) did not show significant differences in their currents compared with control cells (Figure 2A blue line). Addition of ionomycin (IO) did not induce a significant increase in the currents of the control cells, but it elicited a strong current in the TcCAKC expressing cells (Figure 2A red line). Similar activation was observed after incubating the oocytes with thapsigargin to release Ca²⁺ from the endoplasmic reticulum (Figure 2B red line). When extracellular Ca²⁺was chelated by addition of EGTA to the medium, the effect of ionomycin was abolished (Figure 2C), indicating that the activation can be triggered by Ca²⁺ from intracellular stores or by influx from the extracellular media. These results confirm that TcCAKC is able to form functional channels by itself and its activity requires increase in cytosolic Ca²⁺. It is important to point out that all recordings were performed in the presence of 1 μM DIDS to block endogenous currents resulting from the activation of calcium-dependent chloride channels, abundant in X. laevis oocytes (Weber W., 1999). TcCAKC activity induced by thapsigargin (Figure 3A red line) was blocked by 1 mM BaCl₂ (Figure 3A black line) while 4-aminopyridine had no significant effect (Figure 3A blue line). To confirm the selective nature of the channel, we mutated the tyrosine at position 313 for alanine (Y313A) and performed similar experiments as described above. This mutation, located in the middle of the conserved selectivity filter (TVGYG), is predicted to render an inactive channel (Heginbotham et al., 1994; Noskov and Roux, 2006). Indeed, oocytes expressing TcCAKC-Y313A showed no current activation upon treatment with ionomycin (Figure 3B green line).

Reversal potential is the voltage at which there is no net current across the membrane and is indicative of the type of ions permeating through a membrane (Hille, 1978). Voltage ramps between −80 and 40 mV were used to test if TcCAKC activation leads to differences in oocyte permeability to K⁺. TcCAKC expressing oocytes treated with ionomycin had a shift in reversal potential of −19.6 mV (Figure 3C red), while control and TcCAKC-Y313A expressing oocytes had almost no shifts in reversal potential [Figure 3C control −2.14 mV (black) and TcCAKC-Y313A 0.43 mV (green)]. The shift in reversal potential for the TcCAKC oocytes is in the direction of the theoretical Nernst Potential for K⁺ (−80.5 mV, assuming [K⁺]_intra of the oocyte is 120 mM, based on previous literature; Weber W. M., 1999), indicates an increase in membrane permeability to K⁺, further confirming TcCAKC as a K⁺ conducting channel.

Overall, the functional complementation of K⁺ deficient yeast and the biophysical characterization of TcCAKC expressed in X. laevis oocytes provide solid evidence that this is a calcium-activated potassium channel that can form functional pores without co-expression of additional subunits.

Since other CAKCs have major influences on membrane potential modulation in eukaryotic cells, it was of interest to elucidate its role in T. cruzi. To test this, fluorometric measurements of resting membrane potential and membrane potential responses were performed to compare differences between WT and mutant epimastigotes. In standard buffer, dKO mutants had a hyperpolarized resting membrane potential (−154.7 ± 5.66 mV, N = 5) when compared to either the WT parasites (−104.1 ± 5.19 mV, N = 5) and sKO parasites (−97.03 ± 9.63 mV, N = 5) (Table 1, Figure 5A). This result suggests that TcCAKC plays a role in membrane potential maintenance in T. cruzi by allowing the influx of K⁺ to the cells. These results are in agreement with previous research, showing that K⁺ causes depolarization on epimastigotes and suggesting the presence of a K⁺ conducting pathway (Van Der Heyden and Docampo, 2002) responsible for this effect.

The WT and knockout mutants were recorded to ion depleted buffers to test if the extracellular ionic composition has an effect on membrane potential in the parasite. Na⁺ free, and K⁺ free buffers (Table 1) did not have significant effects on the resting membrane potential of any of the cell lines. Replacement of Na⁺ and K⁺ with non-permeating cation N-methyl-D-glucamine (NMDG), produced significant depolarization in WT (−120.40 ± 7.95 mV) and sKO (−125.00 ±7.45 mV) parasites but did not further hyperpolarize the dKO epimastigotes (Figure 5B). These results support previously reported evidence that T. cruzi membrane potential is not maintained by Na⁺/K⁺ balance, but is instead primarily dependent on protons (Van Der Heyden and Docampo, 2000). Our results also indicate that, although Na⁺ and K⁺ are not the main driver of membrane potential, the presence of at least one of these ions is required for proper membrane potential homeostasis, presumably to fuel exchangers that contribute to the proton-motive force. To maintain the electrochemical gradients, TcCAKC could be one of the K⁺ influx pathways and its ablation in the dKO parasites pushes the membrane potential to more hyperpolarized values.

Since TcCAKC KO had an important effect on membrane potential maintenance, and previous studies have described protons as the primary regulator of membrane potential in T. cruzi (Van Der Heyden and Docampo, 2000), it was vital to interrogate the role TcCAKC had on intracellular pH (pH_i) regulation. The pH_i of WT (7.36 ± 0.131) and sKO (7.45 ± 0.082) epimastigotes was similar to previously reported values (Van Der Heyden and Docampo, 2000) but dKO parasites had a drastic intracellular alkalinization (8.01 ± 0.124) (Figure 6A solid bars, Table 2). No ionic replacement (Na⁺. K⁺, or NMDG) caused a significant change in the cytosolic pH of the WT parasites, but replacement of both Na⁺ and K⁺ with NMDG caused acidification in sKO and dKO epimastigotes suggesting a less robust homeostatic potential when TcCAKC expression is decreased (Figure 6A, striped bars and Table 2). These results confirm previous findings in T. cruzi showing an interdependency of pH and membrane potential, linked via proton regulation (Van Der Heyden and Docampo, 2002; Vieira et al., 2005). As TcCAKC dKO cells present a hyperpolarized membrane potential, this is also transduced in a relative proton deficit and the observed cytosolic alkalinization.

To further explore the link between TcCAKC-mediated K⁺ influx and pH regulation we performed pH_i measurements in standard buffer in the presence of 1 mM BaCl₂, a blocker that showed a strong effect in the TcCAKC currents in oocytes (Figure 3A). This divalent cation induced a significant acidification in the WT parasites (Figure 6B) but only a marginal effect in the dKO, indicating that the K⁺ influx through the channel is necessary for pH compensatory mechanisms, perhaps by K⁺/H⁺ and Na⁺/H⁺ exchangers. To test this hypothesis, we measured the rate of proton extrusion in epimastigotes and found a significantly higher rate of proton extrusion in dKO parasites (Figure 6C red line) compared with WT (Figure 6C black line). Rate of extrusion measured in extracellular buffer lacking K⁺ was reduced by 34% both in WT and dKO parasites (Figure 6D white bars) and was almost completely eliminated in absence of Na⁺ (Figure 6D gray bars) or when both ions were replaced by NMDG (Figure 6D striped bars). This provides strong evidence of the presence of active ion exchangers in T. cruzi.

Calcium plays a central role in T. cruzi, regulating cell infectivity (Moreno et al., 1994; Caradonna and Burleigh, 2011) and signaling (Burleigh and Woolsey, 2002; Docampo and Huang, 2014). Intracellular calcium balance depends on strict membrane potential regulation as the main plasma membrane permeation pathways are voltage-gated calcium channel (Verheugen et al., 1995; Christel and Lee, 2012; Harraz and Altier, 2014). Thus, we investigated whether TcCAKC function affects calcium homeostasis in the parasites. Fluo2-AM loaded dKO epimastigotes in BAG had a lower steady state intracellular calcium (55.4 ± 8.84 nM) compared with WT (105 ± 6.69 nM) and sKO (98.5 ± 9.05 nM) as it is shown in Figures 7A,B. Upon addition of 1.8 mM extracellular calcium, WT and sKO show a robust increase in cytosolic calcium (Figure 7A black and blue lines, respectively), while dKO have a modest increase (Figure 7A red line), reaching levels similar to WT under baseline conditions (first 50 s). Given the significant difference in intracellular calcium observed in the TcCAKC dKOs, we compared calcium concentrations under ionic replacement conditions by normalizing the values respect to the initial fluorescence ratio for each cell line. WT epimastigotes show a reduced cytosolic calcium increase when under Na⁺ free, K⁺ free or NMDG conditions (Figure 7C and Table 3) indicating that, at least 60% of the increase is dependent of monovalent cations. This effect can be attributed to direct activity of channel and exchangers or indirectly through decrease of the open probability of voltage-gated calcium channels, as we have observed that in absence of monovalent cations epimastigotes are hyperpolarized (Figure 5B). As expected, the cytosolic calcium levels stayed lower in dKOs under all ionic conditions (Figures 7D,E and Table 3).

In summary, ablation of TcCAKC has a profound effect on cellular homeostasis, with epimastigotes showing a significant hyperpolarization, increase in intracellular pH and rate of proton extrusion, and decrease in cytosolic Ca²⁺concentrations. These results, together with the observed reduction in growth rate and the inability to produce sustained infection in mammalian cells supports the role of TcCAKC as key regulator of T. cruzi physiological fitness.