Functional crosstalk between the TIM22 complex and YME1 machinery maintains mitochondrial proteostasis and integrity

Mitochondria are often referred to as the powerhouse of the cell due to their ability to generate ATP. However, apart from energy conversion, this organelle is also involved in diverse cellular processes, including iron–sulfur cluster production, biosynthesis of several metabolites and apoptosis (Lill and Mühlenhoff, 2005; Nunnari and Suomalainen, 2012; Kang et al., 2018). Therefore, proper mitochondrial function is paramount for cell survival, and any impairment in mitochondrial health could be detrimental to cells. Mitochondria encompass a wide variety of proteins to execute these essential cellular functions, most of which are encoded by the nuclear genome, translated in the cytosol, and subsequently translocated into different mitochondrial subcompartments with the help of specialized protein-conducting machinery known as translocases (Baker et al., 2007; Neupert and Herrmann, 2007; Endo and Yamano, 2009; Wiedemann and Pfanner, 2017). The translocase of the outer membrane (TOM) complex functions as a general entry gate for most incoming mitochondrial proteins (Dukanovic and Rapaport, 2011; Araiso et al., 2019). On the contrary, the inner membrane (IM) contains two translocases: the TIM23 complex (presequence translocase) and the TIM22 complex (carrier translocase) (Chacinska et al., 2009; Schmidt et al., 2010; Dudek et al., 2013). The TIM23 complex mediates the sorting of proteins with presequences into the IM and the matrix, whereas the TIM22 complex integrates polytopic membrane proteins into the IM (Sirrenberg et al., 1996; Rehling et al., 2004; van der Laan et al., 2010; Schendzielorz et al., 2018; Kumar et al., 2022).

The TIM22 complex is evolutionarily conserved across the eukaryotic system, but its composition differs from yeast to humans. In yeast, the TIM22 complex is a 300-kDa insertase machinery consisting of the membrane subunits Tim22, Tim54, Sdh3 and Tim18, and small Tim proteins Tim9, Tim10 and Tim12 (Jarosch et al., 1996, 1997; Kerscher et al., 1997, 2000; Adam et al., 1999; Koehler et al., 2000; Kovermann et al., 2002; Gebert et al., 2011; Stojanovski et al., 2012). Tim22 forms the central core channel, and its conserved regions play a critical role in recruiting the partner subunits to assemble the functional translocase (Rehling et al., 2003; Kumar et al., 2020; Zhang et al., 2021). On the other hand, Tim18 and Sdh3 provide stability to the TIM22 complex, whereas Tim54 acts as an adaptor protein for docking of small Tim chaperones (Kerscher et al., 2000; Koehler et al., 2000; Wagner et al., 2008; Gebert et al., 2011). The small Tim proteins assist in transferring hydrophobic client proteins from the TOM complex to the TIM22 complex (Wagner et al., 2008; Weinhäupl et al., 2021). The TIM22 pathway consists of an additional hexameric complex formed by another class of small Tim proteins, Tim8 and Tim13, required for the biogenesis of certain specific substrates, such as Tim23 (Leuenberger et al., 1999; Koehler et al., 1999). On the contrary, the human TIM22 complex is 440-kDa protein machinery containing the channel-forming module Tim22 and small Tim chaperones Tim9, Tim10a and Tim10b (functional homolog of yeast Tim12) (Bauer et al., 1999; Mühlenbein et al., 2004; Qi et al., 2021). Furthermore, two additional proteins, Tim29 and AGK, have been identified as novel membrane subunits of the human TIM22 complex (Callegari et al., 2016; Kang et al., 2016, 2017; Vukotic et al., 2017). Despite having subtle changes in the structural composition, the TIM22 complex and its import pathway remain conserved from yeast to higher eukaryotes (Kumar et al., 2022). Importantly, pathological variants in different subunits of the TIM22 complex are identified and implicated in various diseases such as mitochondrial myopathy, Sengers syndrome and Mohr–Tranebjaerg syndrome (deafness–dystonia–optic neuropathy syndrome) (Koehler et al., 1999; Roesch et al., 2002; Kang et al., 2017; Vukotic et al., 2017; Pacheu-Grau et al., 2018). These studies highlight the role of defective mitochondrial protein biogenesis through the TIM22 pathway in neurological and musculoskeletal disease development.

The model substrates of the TIM22 pathway include the large family of metabolite carrier proteins such as Pic (also referred to as Pic2), Aac2 (also referred to as Pet9) and Dic1, which all have six transmembrane (TM) segments (Endres et al., 1999; Horten et al., 2020). The TIM22 complex also mediates the membrane insertion of specific components of the translocases (Tim17, Tim22 and Tim23) that contain four TM regions (Leuenberger et al., 1999; Horten et al., 2020; Paschen et al., 2000). Recently, mitochondrial pyruvate carriers (MPCs), which possess three TM segments, and sideroflexins (SFXNs), which have five TM regions, were identified as unconventional client proteins of the TIM22 pathway (Gomkale et al., 2020; Rampelt et al., 2020; Acoba et al., 2021; Jackson et al., 2021). The substrates of the TIM22 pathway are highly hydrophobic, and their proper assembly is critical for maintaining mitochondrial proteostasis. Thus, the turnover of these protein substrates needs to be efficient in maintaining mitochondrial homeostasis.

Mitochondrial protein quality control is maintained by multiple mechanisms such as mitochondrial dynamics, mitophagy, proteasome-assisted protein degradation, chaperones and proteases (Ruan et al., 2020; Song et al., 2021). Amongst them, mitochondrial proteases operate at the molecular level and regulate quality control, ensuring a healthy mitochondrial pool (Koppen and Langer, 2007). Yme1 is an IM mitochondrial protease involved in the degradation of misfolded or unfolded IM or intermembrane space proteins, and specific outer membrane (OM) proteins (Leonhard et al., 2000; Gerdes et al., 2012; Wu et al., 2018). It belongs to the i-AAA family of proteases that function in an ATP-dependent manner (Leonhard et al., 1999). Structurally, the YME1 complex encompasses the core component Yme1 and two additional subunits, Mgr1 and Mgr3 (Dunn et al., 2008). Yme1 is a zinc metalloprotease consisting of a protease domain and an ATPase domain that work together for substrate processing, whereas Mgr1 and Mgr3 act as adaptor proteins (Puchades et al., 2017). Yme1 regulates multiple arrays of the proteins, subunits of the electron transport chain such as Cox2 and Cox4, small Tim protein Tim10 and mitophagy-selective receptor Atg32 (Weber et al., 1996; Stiburek et al., 2012; Wang et al., 2013; Spiller et al., 2015). Moreover, the loss of Yme1 results in abnormal mitochondrial morphology and increased frequency of mitochondrial DNA escape (Thorsness et al., 1993; Campbell and Thorsness, 1998; Cesnekova et al., 2018).

In higher eukaryotes, YME1L1, the functional homolog of Yme1, is involved in the cleavage of OPA1, a key regulator of mitochondrial fusion and fission (Griparic et al., 2007; Tilokani et al., 2018). Furthermore, mutations in YME1L1 are associated with optic atrophy (Hartmann et al., 2016). Additionally, Tim17A, a substrate of the TIM22 pathway, was shown to be the degradation substrate of YME1L1, suggesting a possible interplay between quality control and the mitochondrial IM import machinery (Rainbolt et al., 2013). Interestingly, in yeast, Tim54, a membrane subunit of the TIM22 complex, is required to assemble Yme1 into the active proteolytic complex (Hwang et al., 2007). Furthermore, Yme1 was demonstrated to be required for quality control of Aac2 (Liu et al., 2015). Taken together, these findings suggest a possible interplay between the mitochondrial quality and IM import machinery via functional crosstalk between the TIM22 complex and YME1 machinery.

The present study demonstrates a novel functional association between the TIM22 complex and YME1 machinery in Saccharomyces cerevisiae. Our genetic, biochemical and cell biological analyses provide compelling evidence to highlight the functional dependence of the TIM22 complex and YME1 machinery in regulating IM protein quality control and, thus, overall mitochondrial health.

To understand the significance of the TIM22 complex in mitochondrial quality control, we investigated the effect of the impaired TIM22 translocase activity in cells devoid of Yme1. To begin with, we generated single-deletion strains, tim18Δ (impaired TIM22 complex) and yme1Δ, and a double-deletion strain tim18Δ yme1Δ in the W303 haploid background. Upon in vivo phenotypic analysis, the tim18Δ strain showed a similar growth pattern as the wild type (WT) at all the indicated temperatures in yeast extract peptone dextrose (YPD) and yeast extract peptone glycerol (YPG) media (Fig. 1A). On the contrary, the yme1Δ strain exhibited severe growth defects at 37°C in YPG and a mild growth sensitivity at 14°C in YPD and YPG media, in agreement with previous reports (Fig. 1A) (Thorsness et al., 1993; Thorsness and Fox, 1993). The disruption of both Tim18 and Yme1 led to slight growth defects at 37°C in YPD medium and 14°C in both YPD and YPG media (Fig. 1A). Strikingly, the tim18Δ yme1Δ strain displayed better restoration of cellular growth compared to yme1Δ cells at 37°C in YPG medium (Fig. 1A). Additionally, when we performed growth-curve analysis at 37°C in YPG medium, the tim18Δ yme1Δ strain showed a significant growth rescue compared to yme1Δ cells (Fig. 1B). The identity of the deletion strains was further validated by immunoblotting analysis (Fig. 1C). Moreover, the complementation of the tim18Δ yme1Δ strain with a construct expressing Tim18 under its native promoter displayed a severe growth sensitivity at 37°C in non-fermentable medium [synthetic complete glycerol (SCG)–Ura], comparable to yme1Δ cells (Fig. 1D). Tim18 protein expression in the tim18Δ yme1Δ strain was confirmed by immunoblotting (Fig. 1E). Furthermore, we used tim22 conditional mutants to validate the genetic link between these complexes (Kumar et al., 2020). Interestingly, the deletion of Yme1 in a tim22 mutant background (tim22^K127A) exhibited partial rescue in the growth defects of yme1Δ at 37°C in YPG medium (Fig. 1F). Taken together, these findings provide genetic evidence for a novel functional link between the TIM22 and YME1 complexes.

Next, we examined whether the rescue of growth defects of yme1Δ is specific to Tim18 loss or a consequence of impairment of the TIM22 complex. To address this, Tim18 and Tim22 were overexpressed under the TEF promoter and we analyzed their effect on the growth of WT and yme1Δ strains by serial dilution assays. Remarkably, the overexpression of Tim22 compromised the growth of cells lacking Yme1 at all tested temperatures in both fermentable [synthetic complete dextrose (SCD)–Ura] and non-fermentable (SCG–Ura) media (Fig. S1A). At the same time, the overexpression of Tim22 did not affect the growth of the control WT strain (Fig. S1A). In contrast, the overexpression of Tim18 did not alter the growth phenotypes of either the WT or yme1Δ strains (Fig. S1B). Tim22 and Tim18 protein overexpression were confirmed by immunoblotting (Fig. S1C,D). In conclusion, these results establish that impairment in the TIM22 complex suppresses the respiratory growth defect of cells devoid of Yme1 at elevated temperatures, highlighting an intricate genetic link between the carrier translocase machinery and the YME1 complex.

To gain insights into the respiratory growth defects of cells lacking Yme1, we first examined the oxygen consumption rate (OCR) in WT and deletion strains at 37°C in YPG medium using a Seahorse XF HS mini analyzer. Upon measurement, WT and tim18Δ cells did not show any significant change in the basal respiration rates (Fig. 2A; Fig. S2A). However, the basal OCR was drastically reduced in yme1Δ strains, suggesting that loss of Yme1 makes cells incompetent (Fig. 2A; Fig. S2B). Remarkably, the basal OCR was rescued in the tim18Δ yme1Δ strains, indicating that loss of Tim18 mitigates the respiratory incompetency of cells devoid of Yme1 (Fig. 2A; Fig. S2C). Additionally, we analyzed the maximal respiration capacity of the WT and deletion strains after treatment with carbonyl cyanide-p-trifluoromethoxyphenylhydrazone (FCCP), a mitochondrial oxidative phosphorylation uncoupler. Upon quantification, we observed that WT and tim18Δ cells exhibited comparable maximal respiration rates (Fig. 2B; Fig. S2A). Conversely, yme1Δ strains showed severe impairment in the maximum respiration capacity (Fig. 2B; Fig. S2B). Notably, the maximal respiration rate was significantly rescued in tim18Δ yme1Δ cells (Fig. 2B; Fig. S2C).

We next analyzed mitochondrial structures, as defective respiration is an indication of impaired mitochondrial integrity. As the deletion of Tim18 rescued the growth of yme1Δ cells (tim18Δ yme1Δ) in respiratory medium, we hypothesized that a possible reversal of mitochondrial integrity could occur through productive, functional crosstalk between the TIM22 and YME1 complexes. To validate this hypothesis, the mitochondrial morphology of WT, tim18Δ, yme1Δ and tim18Δ yme1Δ strains was examined by microscopy. The mitochondrial network was visualized using the MTS–mCherry construct, which exclusively decorates mitochondria. WT and deletion strains expressing MTS–mCherry were grown in glycerol-containing medium at the permissive temperature, and mitochondrial morphology was monitored by fluorescence imaging. Most of the WT and tim18Δ cells displayed a reticular or tubular mitochondrial network (Fig. 2C,D). However, the yme1Δ strain demonstrated fragmented or punctate mitochondrial structures (Fig. 2C,D), in agreement with previous findings (Campbell and Thorsness, 1998; Campbell et al., 1994). Interestingly, tim18Δ yme1Δ cells showed enrichment of reticular or tubular mitochondrial networks, suggesting that the deletion of Tim18 can rescue the mitochondrial morphological defects associated with yme1Δ cells under respiratory growth conditions (Fig. 2C,D). Taken together, these results suggest that functional interplay between the TIM22 and YME1 complexes is crucial for mitochondrial integrity and cellular respiration.

To further delineate the importance of the genetic link between the TIM22 and YME1 complexes in mitochondrial health, we examined different mitochondrial functional parameters. YME1 was identified in a genetic screen for isolating genes that manifest an increased frequency of mitochondrial DNA migration to the nucleus (Thorsness and Fox, 1993). We assessed mitochondrial DNA stability as one of the parameters to evaluate mitochondrial health. We used a genetic approach to examine where the nuclear TRP1 gene, including its regulatory regions, was integrated into the mitochondrial genome in a strain lacking its endogenous TRP1 gene (Fig. 3A) (Thorsness et al., 1993). The mitochondrial TRP1 gene, when migrated to the nucleus, becomes functional as the required transcription machinery is only present in the nucleus. The presence of TRP1 in mitochondrial DNA permits monitoring of the rate at which mitochondrial DNA fragments escape from the organelle. Upon assessment, the yme1Δ strain demonstrated higher mitochondrial DNA transfer frequency in a time-dependent manner consistent with earlier findings (Fig. 3B,C) (Thorsness et al., 1993; Cheng and Ivessa, 2010). However, the tim18Δ yme1Δ strain showed mitochondrial DNA transfer frequency comparable to the WT and tim18Δ strains, indicating that loss of Tim18 reduces the mitochondrial DNA escape frequency of cells devoid of Yme1 (Fig. 3B,C). As a loading control, an equal number of cells from the WT and deletion strains were serially diluted and spotted on YPD medium (Fig. 3D). To further corroborate the above result, we checked for mitochondrial DNA content using the mitochondrial DNA-specific dye SYTO 18. The WT and deletion strains were grown in YPG medium under permissive conditions, followed by staining with SYTO 18 and visualization by fluorescence imaging. Strikingly, the yme1Δ strain displayed reduced levels of SYTO 18 fluorescence than WT and tim18Δ strains (Fig. 3E). In contrast, the tim18Δ yme1Δ strain exhibited SYTO 18 fluorescence levels comparable to those of the WT and tim18Δ strains, implying that the deletion of Tim18 decreases the mitochondrial DNA loss frequency of cells deficient in Yme1 (Fig. 3E).

We measured the ATP levels as another criterion for mitochondrial function in deletion strains to validate the above results. Upon evaluation, the levels of mitochondrial ATP were drastically reduced in yme1Δ cells compared to WT and tim18Δ cells (Fig. 3F). On the contrary, the deletion of Tim18 partially restored the ATP levels in cells lacking Yme1, indicating that the growth defects of the yme1Δ cells in respiratory medium are a consequence of the impairment in mitochondrial functions (Fig. 3F). Porin (Por1), an OM protein, was used as a control to ensure that equal amounts of mitochondria were used for the analysis (Fig. 3F). Collectively, these findings signify that the functional link between the TIM22 complex and YME1 machinery plays an essential role in conserving mitochondrial functions.

To get mechanistic insights into the significance of the association between the TIM22 complex and YME1 machinery, we analyzed the steady-state protein levels of TIM22 pathway substrates in the cellular lysates of the WT and deletion strains grown in YPG medium at the permissive temperature. The protein levels of Dic1, Aac2 and Tim23 were observed to be similar between WT and tim18Δ strains (Fig. 4A,B). However, the yme1Δ strain showed higher levels of Dic1, Aac2 and Tim23 than the WT strain (Fig. 4A,B). Remarkably, the levels of these proteins were significantly reduced in the tim18Δ yme1Δ strain compared to the yme1Δ strain (Fig. 4A,B). As controls, the levels of Por1 and PGK1 remained unaltered in the WT and all the deletion strains (Fig. 4A,B). To delineate the functional coordination of the TIM22 pathway and YME1 machinery at the organellar level, the steady-state expression of TIM22 complex substrates was analyzed from isolated mitochondrial lysates. Interestingly, the mitochondrial levels of Dic1, Aac2 and Tim23 were comparable between WT and all the deletion strains, indicating that the absence of Yme1 results in the accumulation of these proteins outside the mitochondria (Fig. 4C,D).

We next checked whether the growth impairment of yme1Δ cells was due to the enhanced levels of TIM22 substrates, in which case overexpression of these cargos would aggravate the growth sensitivity of yme1Δ cells. To test the hypothesis, metabolite carrier proteins (Pic, Aac2 and Dic1) and Tim23 were overexpressed, followed by growth phenotype analysis. The overexpression of Dic1 did not affect the growth of WT and tim18Δ cells compared to the yme1Δ strain, which exhibited severe growth defects at all temperatures in fermentable medium (SCD–Trp) (Fig. 5A). Additionally, tim18Δ yme1Δ cells showed severe growth defects upon Dic1 overexpression under the same experimental conditions (Fig. 5A). This signifies that the activity of carrier translocase and protease systems is crucial for handling the excess TIM22 pathway substrates. The overexpression of Dic1 was confirmed by immunoblotting (Fig. 5B). Similarly, the overexpression of Tim23 considerably impaired the growth of the yme1Δ strain at 37°C and further enhanced the growth defects of the tim18Δ yme1Δ strain at all tested temperatures in fermentable medium (SCD–Trp) (Fig. S3A). The overexpressed Tim23 protein levels were validated by immunoblotting in WT and deletion strains (Fig. S3B). Moreover, the overexpression of more complex cargo proteins such as Pic and Aac2 resulted in the lethality of yme1Δ and tim18Δ yme1Δ cells (Fig. S3C,D). As a control, the overexpression of an OM protein, Tom6, or a matrix protein, Abf2, did not alter the growth of either WT or any deletion strains under similar experimental conditions (Fig. S3E,F). Furthermore, the overexpression of Tim18 (an IM protein and substrate of the TIM23 pathway) did not affect the growth phenotype of cells lacking Yme1 (Fig. S1B,D), suggesting that the growth defect of the yme1Δ strain is a consequence of impairment in the TIM22 pathway.

To check whether higher levels of TIM22 pathway substrates influence mitochondrial integrity and thus further impair the cellular growth of yme1Δ cells, we examined the mitochondrial morphology in Dic1-overexpressing cells grown in fermentable medium at the permissive temperature. Intriguingly, WT cells overexpressing Dic1 retained their tubular mitochondrial network, whereas the yme1Δ strain overexpressing Dic1 demonstrated further enhancement in mitochondrial fragmentation even under fermentable growth conditions at the permissive temperature, implying that excess levels of TIM22 pathway substrates impair mitochondrial integrity in the absence of Yme1 (Fig. 5C,D).

A previous finding reported that partial suppression of cytosolic protein synthesis rescues the cellular lethality of yme1Δ cells expressing a pathogenic variant of Aac2 (Wang et al., 2008). To determine whether loss of Tim18 can rescue the growth defects of cells lacking Yme1 following a similar mechanism, we performed the growth assessment of WT, yme1Δ and tim18Δ yme1Δ after cycloheximide (CYH) treatment. Strikingly, CYH treatment did not change the growth pattern of any of the strains compared to untreated cells under our experimental conditions (Fig. 5E). Cytoplasmic protein synthesis inhibition upon CYH treatment was ascertained by performing immunoblot analysis against several cytoplasmic proteins. As indicated in Fig. S4A,B, a reduction in the steady-state levels of cytosolic proteins such as Ydj1, Hsp31 and SOD1 was observed in the deleted strains upon CYH treatment. These results further highlight that the partial suppression of cytoplasmic protein synthesis does not rescue the growth defects of cells lacking Yme1 under respiratory growth conditions. Additionally, the levels of Ssb1 remained unaltered in WT and all the deletion strains at both 30°C and 37°C (Fig. S4C,D). Ssb1 is a cytoplasmic chaperone that regulates the accumulation of proteins in the cytosol and is upregulated in tim18Δ cells lacking mitochondrial DNA (Dunn and Jensen, 2003; Wang and Chen, 2015). Hence, these findings suggest a possible involvement of a novel cellular response associated with the TIM22 pathway and YME1 machinery. In summary, these results suggest that excess protein levels of TIM22 pathway substrates impair mitochondrial integrity, thus leading to the growth sensitivity of cells deprived of Yme1 under respiratory conditions. However, compromising the TIM22 pathway in cells lacking Yme1 preserves mitochondrial morphology and functions, possibly by reducing the levels of TIM22 pathway substrates, leading to cellular growth rescue.

To dissect the functional link between the TIM22 complex and YME1 protease machinery, we next estimated the steady-state protein levels of different components of the carrier translocase machinery and Yme1 in isolated mitochondria from WT and deletion strains grown in non-fermentable medium. Upon analysis, we found a slight reduction in Tim22 protein levels in the mitochondrial lysates of the tim18Δ strain at both 30°C and 37°C compared to those from the WT strain (Fig. S5A; Fig. 6A). This agrees with previous findings in which Tim18 was demonstrated to be involved in maintaining Tim22 stability (Koehler et al., 2000; Kerscher et al., 2000). Furthermore, the levels of Tim22 and Tim18 were marginally reduced in yme1Δ cells at 30°C (Fig. S5A). Strikingly, the levels of Tim22 and Tim18 were significantly reduced together with a minor decline in the levels of Tim54 in cells lacking Yme1 at 37°C (Fig. 6A). Additionally, the tim18Δ yme1Δ strain showed a reduction in Tim22 levels at 30°C, whereas the levels of Tim54 and Tim22 were significantly decreased at 37°C (Fig. S5A; Fig. 6A). On the contrary, the levels of Yme1 protein were comparable between the WT and tim18Δ strains at both 30°C and 37°C (Fig. S5A; Fig. 6A). As positive controls, the amounts of porin and Tim50 (a marker for the IM) were examined and found to be comparable between WT and all the deletion strains (Fig. S5A; Fig. 6A). These results suggest that the loss of Yme1 alters the steady-state levels of different subunits of the TIM22 complex, especially at elevated temperatures.

We further investigated the effect of Yme1 deletion on the stability of the TIM22 complex using Coomassie Blue-stained native PAGE (BN-PAGE). The WT mitochondria isolated from cells grown at 30°C in YPG medium displayed a stable TIM22 complex at ∼300 kDa when immunodecorated with antibodies specific for Tim54 and Tim22 (Fig. 6B). On the contrary, mitochondria isolated from tim18Δ and tim18Δ yme1Δ strains exhibited destabilization of the TIM22 complex (Fig. 6B). This finding agrees with earlier reports in which the loss of Tim18 was shown to impair the architecture of the TIM22 complex (Koehler et al., 2000; Kerscher et al., 2000). Surprisingly, the levels of the 300-kDa TIM22 complex were significantly affected in cells devoid of Yme1 (Fig. 6B). Furthermore, the stability of the TIM22 complex was more drastically affected at 37°C in Yme1-deficient cells (Fig. S5B). This correlates well with the reduction in the levels of Tim22 and Tim18 shown above. On the contrary, the levels of the YME1 complex remained unaffected between the WT and tim18Δ strains, suggesting that loss of Tim18 does not affect the stability of the YME1 complex (Fig. 6B). As a control, the amounts of the porin complex were assessed and found to be comparable between WT and all the deletion strains at 30°C and 37°C (Fig. 6B; Fig. S5B). Taken together, these results highlight that the deletion of Yme1 affects the stability of the TIM22 complex.

To gain more insight into the connection between these systems, we next assessed whether the TIM22 and YME1 complexes physically interact with each other. To test this, we isolated mitochondria from WT cells and strains expressing C-terminally FLAG-tagged Tim18 and subjected them to co-immunoprecipitation (co-IP) using anti-FLAG prebound to protein G sepharose beads. Upon analysis, Tim18–FLAG efficiently immunoprecipitated Tim54 and Sdh3 (Fig. 6C), corroborating with a previous report (Kumar et al., 2020). Intriguingly, we observed co-purification of Yme1 along with Tim18–FLAG (Fig. 6C). Tim44, a subunit of the TIM23 complex, and mitochondrial lysates from WT cells were used as negative controls to determine the specificity (Fig. 6C). Furthermore, we performed a similar co-IP with strains expressing either C-terminally FLAG-tagged Tim23 or SOD1. Interestingly, both Tim23–FLAG and SOD1–FLAG failed to immunoprecipitate Yme1, ruling out the possibility of a FLAG-specific interaction of Yme1 (Fig. 6D,E). Based on these findings, we envisioned that the TIM22 complex and YME1 machinery are in proximity to each other in the mitochondrial IM.

The current study highlights a functional interplay of the TIM22 complex and YME1 protease machinery in regulating mitochondrial health and integrity (Fig. 7). Our genetic analyses indicate that the deletion of Tim18, a membrane component of the TIM22 complex, rescues the respiratory growth defects of cells devoid of Yme1. Additionally, one of the mutations in the TM2 region of Tim22 (tim22^K127A) partially suppresses the growth defects of cells lacking Yme1 under respiratory conditions. Furthermore, by utilizing protein overexpression, biochemical and cell biological studies, we demonstrated that the growth rescue of cells deprived of Yme1 is a consequence of impairment in the TIM22 pathway.

The TIM22 complex is one of the major protein translocases that performs a critical function in the biogenesis of the most intricate polytopic proteins of the mitochondrial IM. However, our understanding regarding the protein homeostasis of the carrier translocase substrates is limited mainly because of a lack of studies related to the regulation of protein turnover under normal and pathological conditions. Therefore, even though the mutations in the various subunits of the TIM22 pathway are known to cause neurological disorders, the underlying mechanisms by which the levels of carrier translocase substrates are regulated under such conditions remain enigmatic.

One of the mechanisms by which cells modulate organelle health is by the action of various proteolytic machinery. Yme1 is an i-AAA mitoprotease that plays a crucial role in mitochondrial quality control by regulating the turnover of several membrane proteins. Although Yme1 is vital for mitochondrial protein turnover, cells have evolved with different adaptative mechanisms to survive in the event of its impairment or absence. Previous studies report mutations in the 26S protease subunit and γ subunit of mitochondrial ATP synthase to repress the mitochondrial function and growth defects of yme1Δ cells without mitochondrial DNA (Campbell et al., 1994; Weber et al., 1995). Another study revealed that partial inhibition of cytosolic protein synthesis rescues the synthetic lethality of cells lacking Yme1 along with a pathogenic variant of Aac2 (Wang et al., 2008). In a similar line, our study highlights that the impairment in the TIM22 complex promoted the survival of Yme1-deficient cells by rescuing mitochondrial health.

The dynamic association of protein translocases with other mitochondrial components plays a crucial role in modulating organelle homeostasis. For example, the TIM22 complex associates with porin (also known as VDAC) and the mitochondrial contact site and cristae-organizing system (MICOS) complex to facilitate carrier protein biogenesis (Callegari et al., 2019; Ellenrieder et al., 2019). On the contrary, the TIM23 complex coordinates with the respiratory complex to maintain the optimum IM potential required for the protein import (Wiedemann et al., 2007; Matta et al., 2020). In agreement, our finding demonstrates similar coordination between the TIM22 complex and YME1 machinery being in proximity to each other to maintain mitochondrial proteostasis and organelle quality control. Notably, the interaction between carrier translocase machinery and the YME1 complex is transient as Yme1 interacts with Tim18–FLAG in a non-stoichiometric manner, as observed in the co-IP. We believe that a small portion of YME1 machinery is associated with the TIM22 complex, giving rise to different pools of protease machinery to manage diverse substrates. At the organelle level, the loss of Yme1 results in punctate mitochondrial structures, enhanced mitochondrial DNA transfer and compromised respiration capacity, which can be suppressed by altering the TIM22 translocase activity. Hence, it is not unreasonable to believe that the IM proteostasis machinery plays a crucial role in the turnover of misassembled proteins in close connection with import machinery.

Previous studies highlight that cells regulate the health of mitochondria by numerous mechanisms, including modulating the components involved in biogenesis and protein translocation rate to avoid the accumulation of misfolded proteins within the organelle. For example, degradation of Tim17A by YME1L1 in higher eukaryotes limits protein import efficiency, thereby assisting mitochondrial proteostasis in response to stress-regulated translation attenuation (Rainbolt et al., 2013). Interestingly, our study provides evidence to show that Yme1 is required for maintaining the stability of the TIM22 complex. We hypothesize that this might be an adaptive strategy to regulate IM protein content due to the unavailability of Yme1. Our study provides compelling evidence that in the absence of Yme1, even though the cells harbored a higher load of substrates due to partial destabilization of the TIM22 complex, the deletion of Tim18 resulted in the relieving the excess of the mitochondrial protein burden, leading to cell survival.

Mitochondrial dysfunction is associated with several pathological states; hence, cells have evolved multiple stress-regulated pathways to maintain mitochondrial proteostasis and organelle function. As the TIM22 pathway substrates are IM proteins with multiple TM segments, any impairment in the TIM22 complex functionality might lead to the accumulation of these highly hydrophobic substrates. Several studies have shown that the accumulation of mitochondrial proteins in the cytosol results in diverse cellular stress responses, such as mitochondrial precursor overaccumulation stress, mitochondrial unfolded protein response, unfolded protein response triggered by mistargeting of proteins, integrated stress response and mitochondrial compromised protein import response (Wang and Chen, 2015; Wrobel et al., 2015; Coyne and Chen, 2018; Weidberg and Amon, 2018). Such stress responses are implicated in several pathological conditions such as aging, muscle degeneration and neurodegeneration (Lin and Beal, 2006; Tatsuta and Langer, 2008; Zhu et al., 2021). In addition, the accumulation of these hydrophobic proteins in the cytosol further dampens the import of other essential cytosolic and mitochondrial proteins (Coyne and Chen, 2019). Additionally, overloading of the carrier proteins in humans induces cytosolic aggresomes and activates apoptotic pathways (Liu et al., 2019). Intriguingly, in our study, yme1Δ cells displayed higher levels of the substrates of the TIM22 pathway outside the mitochondria, possibly in the cytosol. Additionally, the loss of Tim18 in the yme1Δ strain reduced the accumulation of TIM22 pathway cargos, thereby ensuring cell survival. Further studies are required to identify the stress response pathway in the cells lacking Yme1 and Tim18 to gain mechanistic insights into the mitochondrial quality control mediated by the association of these two complexes.

In summary, the present study revealed a unique functional link between the TIM22 pathway and YME1 machinery in maintaining mitochondrial proteostasis and integrity. Thus, our results provide a primary platform to explore the connection between these two complexes in higher eukaryotes which can be crucial in understanding the pathogenesis of several diseases.

The yeast strains used in this study are mentioned in Table S1 (Thorsness et al., 1993, Hines et al., 2011, Okamoto et al., 2014). The details of plasmids and primers used in this study are described in Table S2. The tim18Δ and yme1Δ strains were generated by PCR-mediated homologous recombination to substitute the ORFs with a kanMX4 and hphNT1 selection cassette, respectively, in a W303 haploid background. The tim18Δ yme1Δ strain was created by deleting YME1, as mentioned above, in a tim18Δ strain. The PTY44/tim18Δ and PTY52/tim18Δ strains were made by removing TIM18 in PTY44 and PTY52 strains following a similar approach described above. The WT TIM18 gene from positions −700 to +859 was PCR amplified from W303 yeast genomic DNA for complementation studies. The PCR-amplified products were cloned into the NotI/SalI-digested yeast centromeric vector pRS316 (plasmid gifted by Prof. Elizabeth A. Craig, University of Wisconsin-Madison, USA). For overexpression of Pic, Aac2, Dic1, Tim23, Abf2 and Tom6, the corresponding open reading frames were PCR amplified from yeast genomic DNA and cloned into the pRS414_TEF vector (plasmid gifted by Prof. Elizabeth A. Craig). The TIM22 gene was PCR amplified from yeast genomic DNA and cloned into the pRS416_TEF vector (plasmid gifted by Prof. Elizabeth A. Craig) for overexpression experiments. The Tim22 mutant and other overexpression constructs used in this study are described in detail in an earlier report (Kumar et al., 2020).

Yeast cells were grown in YPD (1% yeast extract, 2% peptone and 2% dextrose), YPG (1% yeast extract, 2% peptone and 2% glycerol), SCD–Trp (0.67% yeast nitrogen base without amino acids, 0.069% Trp dropout supplement and 2% dextrose), SCD–Ura (0.67% yeast nitrogen base without amino acids, 0.072% Ura dropout supplement and 2% dextrose), SCD–Ura–Trp (0.67% yeast nitrogen base without amino acids, 0.069% Ura Trp dropout supplement and 2% dextrose) or SCG–Ura (0.67% yeast nitrogen base without amino acids, 0.072% Ura dropout supplement and 3% glycerol) media.

Strains harboring the kanMX4 or hphNT1 selection marker were selected on YPD supplemented with either 400 μg ml⁻¹ G418 sulfate or 250 μg ml⁻¹ hygromycin. Yeast cells were streaked and cultured on SCD containing 1 mg ml⁻¹ 5-fluoroorotic acid to remove the URA3-harboring plasmid.

For spot assay, the respective yeast strains were grown at 30°C in YPD or the indicated media up to the mid-log phase. Cells were harvested by centrifugation (2200 g for 5 min at room temperature) followed by serial dilution and spotting on YPD, YPG or other required minimal media. The plates were incubated at various temperatures, followed by documentation of images at appropriate time intervals.

For growth curve examination, WT and deletion strains were cultured at 30°C in YPD for 12 h. Following this, an equal amount of cells (absorbance at 600 nm or A₆₀₀=0.07) were harvested by centrifugation (2200 g for 5 min at room temperature), washed twice with sterile water and re-inoculated in 50 ml YPG. Absorbance was measured at 600 nm every 12 h intervals for 4 to 5 days using an ultraviolet-visible spectrophotometer (Eppendorf, Hamburg, Germany).

Total cell protein lysates were prepared by the trichloroacetic acid (TCA; Sigma-Aldrich, T0699-100ML) precipitation method. Equal numbers of cells (A₆₀₀=5) were harvested by centrifugation at 16,900 g for 5 min at room temperature, resuspended in 10% TCA, vortexed thoroughly and incubated for 30 min at 4°C with gentle vortexing at 10 min intervals. The samples were centrifuged (2200 g for 5 min at 4°C) and TCA was removed completely. The resultant pellet was washed twice with acetone, followed by incubation at 42°C for 10 min. Acid-washed 0.5 mm glass beads were added in a 1:1 ratio (w/w) to the pellet along with 1× SDS-PAGE sample buffer (50 mM Tris-HCl, pH 6.8, 2% SDS, 0.1% Bromophenol Blue, 10% glycerol and 100 mM β-mercaptoethanol) and vortexed at maximum speed ten times with intermittent incubation at room temperature. The samples were boiled at 90°C for 10 min, and the final whole-cell lysate was extracted by centrifugation (16,900 g for 10 min at room temperature). Lysate volumes equivalent to A₆₀₀=0.5 or 1 (depending on protein abundance) were resolved by SDS-PAGE followed by western blotting.

The respiratory capacity was assessed using the Seahorse XF HS mini analyzer (Agilent Technologies, Santa Clara, CA). Seahorse XFp cell culture miniplates were coated with 50 μg ml⁻¹ poly-D-lysine (50 μl each well) (Sigma-Aldrich, P6407-5MG) for 30 min at room temperature, and then aspirated followed by air drying at room temperature for 2 h. The Seahorse XFp extracellular flux cartridge was hydrated with sterile water (200 μl per well) and incubated overnight at 37°C in a non-CO₂ incubator and Seahorse XF calibrant solution (Agilent Technologies). After incubation, the water was removed, then the calibrant solution (200 μl per well) was added to the sensor cartridge and incubated at 37°C for 1 h before measurement.

Yeast cultures of WT and the deletion strains were grown in YPD medium at 30°C for 12 h, followed by subculturing in YPG medium at 30°C for 8 h. The cultures were then shifted to 37°C for 8 h to induce the phenotype. For measurement, all cultures were diluted to A₆₀₀=0.01 to seed cells in wells into a poly-D-lysine-coated Seahorse XFp cell culture miniplate in 50 μl of Seahorse XF assay medium (Agilent Technologies). For background measurement, two wells containing only assay medium were included. The loaded plate was centrifuged at 300 g for 1 min at room temperature with no brakes. After centrifugation, the volume of the medium was made up to 180 μl, and the loaded plate was incubated for 30 min at 37°C in a non-CO₂ to facilitate the transition of the plate into the temperature of the Seahorse machine. For maximal and non-mitochondrial respiration rates, 4 μM FCCP was added to port A (Seahorse XFp Cell Mito Stress Test Kit, Agilent Technologies, Wilmington, DE) and 1.4 μM rotenone/antimycin A in port B (Agilent Seahorse XFp Cell Mito Stress Test Kit). Four readings were taken for basal respiration, whereas six measurements were recorded after FCCP and rotenone/antimycin A injections.

The mitochondrial morphology was analyzed following a prior procedure (Bankapalli et al., 2015). Yeast strains were transformed with the pRS416_TEF MTS–mCherry construct (Bankapalli et al., 2015) to visualize the mitochondrial morphology. MTS–mCherry specifically decorates the mitochondria as it contains MTS from subunit 9 of ATP synthase (pSU9), thereby allowing visualization of the mitochondrial network. WT and deletion strains were grown to the mid-log phase in SCG–Ura at 30°C. The cells were then harvested by centrifugation (2200 g for 5 min at room temperature) followed by final suspension in 1× PBS. The slides were prepared by placing cells (A₆₀₀=0.1) on 2% agarose pads and covering them with a coverslip. The images were acquired with a Delta Vision Elite fluorescence microscope (GE Healthcare) using a 100× objective lens by keeping a uniform exposure throughout the experiment. The excitation wavelength of 587 nm and the emission wavelength of 610 nm were used for mCherry fluorescence imaging. The images were deconvolved and examined using SoftWoRx 6.1.3 software (GE Healthcare).

For mitochondrial DNA visualization, the corresponding yeast cells were allowed to grow up to the mid-log phase in YPG at 30°C and harvested by centrifugation (2200 g for 5 min at room temperature). The cells were resuspended in 1× PBS followed by staining with 10 µM SYTO 18 [Yeast Mitochondrial Stain Sampler Kit (Cat. No. Y7530), Thermo Fisher Scientific] for 15 min at 30°C. After this, the cells were thoroughly washed twice with 1× PBS and placed on agarose pads. Images were captured with Delta Vision Elite Fluorescence Microscope (GE Healthcare) using a 100× objective lens. The excitation wavelength of 468 nm and the emission wavelength of 533 nm was used for monitoring SYTO 18 fluorescence. The images were processed using SoftWoRx 6.1.3. software.

The rates of mitochondrial DNA transfer were determined as described previously (Thorsness et al., 1993; Thorsness and Fox, 1993). Yeast strains were grown in YPD and equal numbers of cells (A₆₀₀=0.5) were harvested by centrifugation (2200 g for 5 min at room temperature). The cells were resuspended in sterile water, followed by plating on a synthetic growth medium lacking the amino acid tryptophan. The number of colonies obtained was counted to calculate the frequency of mitochondrial DNA transmission.

ATP levels were estimated using the ATP detection reagent of the Mitochondrial ToxGlo Assay kit (Promega, Madison, WI). 50 μg mitochondria from WT and deletion strains grown at 30°C in YPG medium were lysed by snap freezing the mitochondria in liquid nitrogen followed by thawing on ice twice. The ATP detection substrate was added in a 1:1 volume to the mitochondrial lysate, and the luminescence was recorded using Glomax Explorer (Promega).

BN-PAGE was carried out following a previously published protocol (Wittig et al., 2006). Briefly, mitochondria (2 mg ml⁻¹) were solubilized in 100 µl of digitonin buffer (1% digitonin, 50 mM NaCl, 50 mM imidazole, 2 mM 6-aminohexanoic acid, 1 mM EDTA, pH 7.0) for 30 min at 4°C. The solubilized portion was separated by centrifugation (16,900 g for 20 min at 4°C), and 15 µl of sample buffer (2.5 µl of 5% Coomassie Brilliant Blue G and 12.5 µl of 50% glycerol) was added per 100 µl of the supernatant fraction. Samples were run on a 6–16% gradient native imidazole PAGE gel, followed by western blotting and immunodetection using antibodies specific to Tim54, Tim22 and porin.

Mitochondria (5 mg ml⁻¹) were solubilized in 1 ml of digitonin buffer [1% digitonin, 25 mM Tris-HCl pH 7.4, 50 mM KCl, 5 mM EDTA, 10% glycerol, 1 mM phenylmethylsulfonyl fluoride (PMSF)]. The insoluble portion was removed by centrifugation (16,900 g for 20 min at 4°C), and the solubilized fraction was then incubated with 100 µl of protein G Sepharose beads prebound to anti-FLAG antibodies. The samples were allowed to gently rotate at 4°C for 12 h followed by three washes with 1 ml of buffer (0.05% digitonin, 25 mM Tris-HCl, 50 mM KCl, 5 mM EDTA, 10% glycerol, 1 mM PMSF, pH 7.4). The immunoprecipitated proteins were eluted by boiling at 65°C for 10 min in a 1× SDS-PAGE sample buffer. The samples were separated on SDS-PAGE and examined by immunoblotting using specific antibodies for the indicated proteins.

Antisera specific for Tim22 (1:250), Aac2 (1:250) and Dic1 (1:250) were gifted by Prof. Agnieszka Chacinska (Warsaw University, Poland). Antisera against Sdh3 (1:250) and Yme1 (1:250) were gifted by Prof. Nikolaus Pfanner (University of Freiburg, Germany). Antiserum specific for Tim54 (1:1000) was gifted by Prof. Toshiya Endo (Kyoto Sangyo University, Japan). Antisera against Tim44 (1:5000), Ssb1 (1:3000) and Ssa1 (1:3000) were gifted by Prof. Elizabeth A. Craig. Antiserum specific for PGK1 (1:7000) was gifted by Prof. P. N. Rangarajan (Indian Institute of Science, Bangalore, India). Antibodies against Tim18 (1:2500), Hsp31 (1:3000), Tim23 (1:5000) and Tim50 (1:5000) were raised in rabbits as reported previously (Pareek et al., 2013; Bankapalli et al., 2015; Kumar et al., 2020). The anti-FLAG tag antibody (1:1000) raised in mouse (F1804) and anti-SOD1 antibody (1:3000) raised in rabbit (HPA001401) were obtained from Sigma-Aldrich and the anti-porin antibody (1:5000) raised in mouse (16G9E6BC4) was purchased from Invitrogen.

Yeast extract, peptone, dextrose and agar were purchased from BD Difco. Yeast nitrogen base without amino acids (Y2030-01), 5-fluoroorotic acid (207291-81-4), G418 sulphate (108321-42-2), and zymolyase (37340-57-1) were purchased from US Biological Life Sciences. Hygromycin (CAS 31282-04-9) and digitonin (CAS 11024-24-1) were purchased from Calbiochem. Other fine chemicals or reagents were purchased from Sigma-Aldrich unless specified.

Mitochondria were isolated following a previously published protocol (Gambill et al., 1993). Western blotting was performed using standard protocols, and the immunoblots were probed using an enhanced chemiluminescence system from Bio-Rad (1705060). The primers used in this study were manufactured by Sigma-Aldrich, and sequencing reactions were carried out by AgriGenome Labs (India). Uncropped blot images are provided in the Fig. S6.

The protein bands were quantified using ImageJ software. Further calculations were performed in Excel (Microsoft), and all statistical analyses were executed using GraphPad Prism 6.0 software. Significance testing was performed using one-way or two-way ANOVA, and Tukey's multiple-comparison test was used to compare different deletion strain values against WT. P-value of <0.05 was considered significant. All indicated n numbers represent biological replicates.

We thank Prof. Thomas Fox for providing yeast strains (PTY44 and PTY52); Prof. Toshiya Endo (Kyoto Sangyo University, Japan) for giving yeast strains (WT Tim22-316 and Tim18–FLAG/WT Tim22-316) as well as Tim54 antibodies; Prof. Elizabeth A. Craig (University of Wisconsin-Madison, USA) for antibodies against Tim44, Ssb1 and Ssa1 and yeast vectors (pRS316, pRS414_TEF and pRS416_TEF); Prof. Agnieszka Chacinska (Warsaw University, Poland) for antibodies against Tim22, Aac2, and Dic1; Prof. Nikolaus Pfanner (University of Freiburg, Germany) for antibodies against Sdh3 and Yme1; and Prof. P. N. Rangarajan (Indian Institute of Science, Bangalore, India) for antibodies against PGK1. We acknowledge the central facility of Department of Biochemistry for Seahorse-related experiments. We thank Dr. Sandeep M. Eswarappa for providing Glomax Explorer to analyze luminescence-based experiments. We thank Vinaya Vishwanathan of P.D.’s laboratory for their constructive and critical comments related to the manuscript and microscopy experiments.