Tools and techniques to measure mitophagy using fluorescence microscopy

Tools and techniques to measure mitophagy using fluorescence microscopy Nick J Dolman*, Kevin M Chambers, Bhaskar Mandavilli, Robert H Batchelor and Michael S Janes Molecular Probes® Labeling and Detection Technologies, Life Technologies™ Corporation, Eugene, Oregon, USA *Corresponding author. Nick J Dolman Ph.D. Molecular Probes® Labelling and Detection Technologies Life Technologies Corporation Eugene, OR 97402, USA Phone: 541-335-0085 Email: nicholas.dolman@lifetech.com Keywords: mitophagy, autophagy, mitochondria, fluorescence microscopy, GFP, LC3. Word count 5516. Abbreviations: Atg, autophagy-related; CALI, chromophore assisted light inactivation; CCCP, carbonyl cyanide 3-chlorophenylhydrazone; FP, fluorescent protein; FRAP, fluorescence recovery after photobleaching; FRET, fluorescence resonance energy transfer ; GFP, green fluorescent protein; JC-1, 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide; ROS, reactive oxygen species; LAMP1, lysosome-associated membrane protein 1; MAP1LC3, microtubule associated protein 1 light chain 3; PtdIns3K, phosphatidylinositol 3kinase; PtdIns4P 5K, phosphatidylinositol 4-phosphate 5-kinase; PINK1, PTEN induced putative kinase 1;RFP, red fluorescent protein; TMRE, tetramethylrhodamine ethyl ester; TMRM, tetramethylrhodamine methyl ester; TR-FRET, time-resolved fluorescence resonance energy transfer; ULK1, unc-51-like kinase 1; WIPI, WD repeat domain, phosphoinositide interacting 1 Abstract Mitophagy is a specialized form of autophagy that removes damaged mitochondria thereby maintaining efficient cellular metabolism and reducing cellular stress caused by aberrant oxidative bursts. Deficits in mitophagy have been shown to underlie several diseases and a substantial body of research has elucidated key steps in the pathways that lead to and execute autophagic clearance of mitochondria. Many of these studies employ fluorescence microscopy to visualize mitochondrial morphology, mass and functional state. Studies in this area also examine co-localization/recruitment of accessory factors, components of the autophagic machinery and signaling molecules to mitochondria. In this review, we provide a brief summary of the current understanding about the processes involved in mitophagy followed by a discussion of probes commonly employed and important considerations of the methodologies to study and analyze mitophagy using fluorescence microscopy. Representative data, where appropriate, is provided to highlight the use of key probes to monitor mitophagy. The review will conclude with a consideration of new possibilities for mitophagy research and a discussion of recently developed technologies for this emerging area of cell biology. 2 Introduction Cell death can be beneficial to an organism, with well characterized examples including the induction of apoptosis during development.1 However, aberrant cell death is detrimental to an organism through premature loss of cells, especially pronounced when the cells lost are postmitotic cell types such as neurons.2 Alternatively, aberrant cell death may cause inappropriate activation of the immune response to those dead cells.3 Inadequate induction of cell death is also a problem, for example during tumorigenesis.2 Therefore the balance between life and death is both critical and highly regulated through a myriad of cellular signaling pathways.2 Mitochondria mediate cell death through a number of mechanisms. Cell death may be initiated via reactive oxygen species (ROS) production in the mitochondria as well as inflammatory signaling and dedicated cell death pathways involving mitochondria.2 The two major cell death pathways, apoptosis and necrosis, proceed via a loss in mitochondrial membrane potential. Apoptosis and necrosis subsequently display divergent mechanisms following loss of mitochondrial membrane potential and therefore mitochondria act as the last common point in these cell death pathways (see Figure 1).3 Mitochondria can therefore be seen as a key modulator of cell fate, and may set the threshold over which cell death proceeds.4 For this reason, cells possess multiple regulatory and protective mechanisms to shield their mitochondrial pools so as to not predispose them to death. A major mechanism by which a cell can maintain a healthy mitochondrial pool is through clearance of damaged mitochondria via a targeted process known as mitophagy.5 Figure 1 summarizes the role of mitochondrial clearance in maintaining cellular function. A number of environmental factors in the mitochondria, such as increased matrix calcium concentrations, increased reactive oxygen species, and decreased ATP 3 to ADP ratios may lead to mitochondrial stress and subsequent loss of mitochondrial membrane potential.6 The degradation and removal of damaged mitochondria potentially represent an important mechanism to maintain healthy pools of mitochondria and overall cell health (Figure 1). While mitochondria are replenished through division of the existing mitochondrial pool, deficits in this process may shift the equilibrium to a state where the cell contains defective mitochondria, perpetuating inefficient respiratory activity, oxidative stress and damage, and even pro-death signaling in the cell (Figure 1).4,7 The degradation of mitochondria was first described in the 1950’s8 with the term ‘mitophagy’ being coined by John LeMasters9 as a way of tying the removal of damaged mitochondria to the method of bulk clearance of aggregated, excess, damaged and superfluous organelles or proteins by macroautophagy. In yeast, two forms of mitophagy have been categorized: “stress-induced mitophagy” and “maintenance mitophagy”.10 Stress-induced mitophagy results from treatments that directly impact mitochondrial function and serves as a protective mechanism against falling cellular ATP levels and the generation of ROS as mitochondrial function is impaired and oxidative phosphorylation is uncoupled. For a review covering the interplay between ROS and autophagy see ref. 11. Maintenance mitophagy serves to keep a healthy pool of mitochondria by clearing aging mitochondria.10 While the specific signals that initiate autophagic clearance of mitochondria are still being fully described, loss of mitochondrial membrane potential appears to be strongly associated with removal by mitophagy. Mitophagy occurs during normal physiological functioning of cells and examples of its importance for cellular homeostasis are exemplified by the need for mitophagy to occur during 4 reticulocyte maturation.12 Several excellent recent reviews summarize current knowledge on the pathways involved in mitophagy and the relevance of these to disease.5,13-15 Mitochondria exist as a highly dynamic intracellular pool that undergoes constant fission (splitting) and fusion (joining). The balance of these two processes has been shown to regulate mitophagy. The fission of mitochondria acts to promote mitophagy as evidenced by removal of key mediators of fission, such as the FIS116 or DNM1L17 (also called DRP1) that lead to an inhibition of mitophagy. Mitochondrial fusion operates in an opposing manner by diluting damaged mitochondrial components thereby improving the overall health of a mitochondrial pool and inhibiting mitophagy (for a review see ref. 18). Mitochondrial morphology itself can influence the induction of mitophagy. The formation of tubular mitochondrial networks during nutrient deprivation has been shown to inhibit mitophagy via down regulation of DNM1L.19 Furthermore, Shutt et al.20 recently provided an eloquent example of how the essentially pro-survival mechanism of mitochondrial fusion may be transduced in response to cellular stress. In this study, the authors showed that cellular stress, sensed by the key stress regulator, oxidized glutathione, was able to confer mitochondrial fusion through disulphide-mediated mitochondrial fusion oligomers and GTP hydrolysis.20 Mitochondrial membrane potential is a key component in the mitophagy pathway. Twig et al.16 showed that non-fusing mitochondria were more prone to undergo mitophagy and that this pool of mitochondria have a significantly lower mitochondrial membrane potential than the cellular pool of mitochondria that is fusion-competent.16 Mitophagy is commonly preceded by loss of mitochondrial membrane potential, however mitochondrial membrane potential loss alone cannot 5 trigger mitophagy.18 Work from John LeMasters and colleagues using laser-ablation of mitochondria would suggest otherwise as an irreversible mitochondrial membrane potential loss was shown to trigger mitophagy.21 An interesting observation made here was that the targeted degradation of mitochondria by autophagosomes did not show the typical sensitivity to phosphatidylinositol 3-Kinase (PtdIns3k) inhibition displayed by nutrient-deprivation induced autophagy.21 In support of this concept, a recent report showed that, in yeast, mitophagy (but not non-selective forms of autophagy) is dependent on the phosphatidylinositol 4-phosphate 5-kinase (PtdIns4P 5k) PIP5K1B (also called MSS4).22 This argues that mitophagy may proceed via a unique intracellular signaling pathway. A recent report showed that ROS generated from a defined set of mitochondria were sufficient to cause depolarization and that these mitochondria were subsequently degraded by mitophagy.23 This report further highlights the close link between mitochondrial ROS generation, mitochondrial membrane potential and mitophagy. Further work will help to clarify the exact role of mitochondrial membrane potential as a marker and/or signal for the induction of mitophagy. Several studies have provided a description of the events that occur following mitochondrial membrane potential loss (for a review see ref. 24). Briefly, under resting conditions, the PTENinduced putative kinase protein 1 (PINK1) is rapidly degraded on the surface of healthy mitochondria. Upon loss of mitochondrial membrane potential, the proteolysis of PINK1 is impaired thereby causing the accumulation of PINK1 on the surface of damaged mitochondria. PINK1 promotes the recruitment of the E3-ligase PARK2/Parkin via phosphorylation of PARK2 by PINK1. Accumulated and active PARK2 ubiquitinates mitochondria and subsequently recruits SQSTM1/ p62 which binds to the autophagy-specific adaptor protein MAP1LC3/LC3. 6 This process targets mitochondria to autophagosomes and subsequent fusion with the lysosome. This mitochondrial quality control pathway is impaired in inherited forms of Parkinson disease, which is characterized by an accumulation of damaged mitochondria and accounts for dopaminergic neuronal cell death associated with this disease.14 This impairment results through loss of function in the key signaling molecules PINK1 or PARK2.14 The existence of inherited diseases arising from mutations in proteins of the mitochondrial quality control pathway further underline the importance of this pathway to cellular homeostasis. While recruitment of PARK2 and subsequent conjugation of ubiquitin to mitochondria represents a clear mechanism to target mitochondria for autophagic clearance, other mechanisms involving the recruitment of PARK2 have been reported. For example, it has been shown that BNIP3L/NIX1 influences mitochondrial recruitment of PARK2 and can target mitochondria for degradation independent of mitochondrial ubiquitination.25 BNIP3L is able to bind MAP1LC3 through an MAP1LC3-interacting region (LIR) in a similar manner to the LIR-containing protein SQSTM1 and therefore may function as an autophagy receptor for mitochondria.25 Even under basal conditions where a mitochondrion is healthy, BNIP3L is located on the outer mitochondrial membrane and therefore may be able to recruit mitochondria to the autophagosome independent of ubiquitination or may prime mitochondria to be cleared rapidly upon damage.15 A homolog of BNIP3L, BNIP3, localizes to mitochondria, is involved in non-apoptotic cell death, and has been shown to also mediate mitophagy. Like BNIP3L, BNIP3 can bind MAP1LC3 through a putative LIR.15,25 Work in yeast highlighted the protein Atg 32 as a receptor for mitophagy.26,27 Neither BNIP3L nor BNIP1 show homology to Atg32, however they may fulfill a similar role in mammalian cells.15 7 In order to study mitophagy, many processes need to be investigated to gain an accurate measurement of the degree and/or regulation of this process as well as its role in cellular homeostasis and disease. In the following sections, we will discuss tools used to study mitophagy. Specifically, probes that are ideally suited for fluorescence microscopy, exploiting the benefit of spatial resolution afforded with this detection modality. These probes are those that may be used to label mitochondria with simultaneous visualization of the sequestration of mitochondria into autophagosomes and subsequent trafficking to lysosomes, as well as those that report mitochondrial membrane potential. A discussion of the methods used to quantify mitophagy and the image analysis procedures employed will also be provided. Representative data, where appropriate, is shown to highlight the use of key probes to monitor mitochondrial function and/or structure along with autophagic/lysosomal markers. The review will conclude with a consideration of new possibilities for mitophagy research and a discussion of recently developed technologies for this emerging area of cell biology. Monitoring mitophagy through co-localization of mitochondrial probes with markers of the autophagic machinery The role of autophagy in clearing mitochondria has been established through studies that show, under conditions known to damage mitochondria, co-localization of mitochondrial markers with elements of the autophagic machinery. Currently, the majority of studies have used colocalization between mitochondrial markers and the autophagosome-specific marker MAP1LC3. The gene MAP1LC3 is the mammalian homolog of the yeast Atg8 gene although there are other homologs, divided into two families.28 The MAP1LC3 family contains four members: 8 MAP1LC3A, MAP1LC3B, MAP1LC3B2 and MAP1LC3C The second family are the GABAA receptor-associated proteins (GABARAP) which has four members: the first is GABARAP, the second member is GABARAPL1(GABAA receptor-associated protein-like 1)/ATG8L/GEC1(Glandular Epithelial Cell 1), the third member is GABARAPL2 (GABAA receptor-associated protein-like 2)/GATE-16(Golgi-associated ATPase enhancer of 16kDa)/GEF2(ganglioside expression factor 2), with the final member GABARAPL3 (GABAA receptor-associated protein like 3).28 Examples where mitochondria have been labeled along with MAP1LC3 as either an Fluorescent protein (FP) chimera19,21,23,29 or by using antibodies raised against MAP1LC330 are noted. MAP1LC3 reporters can also be combined with many other mitochondrial/mitophagy fluorescent markers, including TMRM,21 mitochondrial-targeted RFP,19 PARK2 as an FP chimera,30,31 PINK1 (Flag epitope-tag PINK1),30 and MitoTracker® dyes.29 MAP1LC3 colocalization with mitochondrial markers provides a reliable indication that the mitochondria in question are destined for autophagic degradation and as as will be discussed later, co-localization of mitochondria with lysosomal markers indicates successful trafficking to the lysosome itself. When examining co-localization with markers of the autophagic machinery, it is important to label mitochondria with as little dependence upon functional state as possible. This can be done with certain specialized fluorescent dyes as well as targeted FPs. The cationic MitoTracker® dyes are commonly used to label overall mitochondrial populations, exploiting the net charge of active mitochondria to indicate the relative degree of mitochondrial polarization at the time of labeling. MitoTracker® dyes carry a thiol-reactive chloromethyl group that covalently binds to 9 proteins within the mitochondria thereby allowing the dye to be better retained within mitochondria.32 For this reason, these dyes can be used to track morphological changes as well as changes in mitochondrial numbers or mass after the induction or inhibition of mitophagy. It should be noted that the MitoTracker® dyes do vary in their propensity to accumulate and to be retained after mitochondrial depolarization. For example Pendergrass et al.33 showed that MitoTracker® Green loading is less dependent on mitochondrial membrane potential than MitoTracker® Red and therefore is better for tracking mitochondrial mass. The retention of any probe should be determined empirically in a given cell type under the experimental conditions used. A similar strategy can be employed to investigate the retention of the dye following loading. Mitochondrial mass and/or structure can also be monitored in live or fixed cells using mitochondrial-targeted FPs. FPs are insensitive to mitochondrial membrane potential and therefore are useful as a counter-label to probes that report mitochondrial function. Figure 2 shows the use of a mitochondrial-targeted RFP in combination with the autophagosomal marker GFP-MAP1LC3B. Under control conditions, GFP-MAP1LC3B is visible throughout the cytoplasm and mitochondria appear as long filamentous structures (Figure 2A I-IV). As cells were cultured in the presence of 10 µM CCCP to depolarize mitochondria and 60 µM chloroquine to block autophagic flux, there was a pronounced accumulation of punctate mitochondrial structures that co-localize with GFP-MAP1LC3B (arrowheads in Figure 2B IV). When using targeted FPs, it is important to consider the effect that gene delivery has upon the cell. This is especially pronounced when studying autophagy.28 It is important that the plasmid 10 DNA isolated is free of contaminants so as to avoid an autophagic response being elicited as a result of entities introduced with the plasmid DNA. Furthermore certain transfection protocols may induce autophagy, including both lipid and viral delivery systems.28 For this reason, it is often beneficial to leave the cells for a period of 48 hours post-transfection to allow equilibration.28 The use of transient expression systems may complicate the interpretation of population-based data as the expression levels are not homogenous across the cells. While the establishment of a stable cell line may be beneficial with respect to this problem, this approach requires a significant investment of time and is not easily transferred from one cell model to another. Stable cell lines are typically generated using immortalized tumor cell lines and are far removed from the physiology of primary human cells and tissues. When using targeted FPs it is important to note that the FP-chimera may affect the biology of the cell. This can occur through either oligomerization of the FP or as a result of over-expression of a targeting moiety (for a review see ref. 34). To gain a measure of changes in endogenous mitochondrial morphology or mass, cells may be labeled with antibodies specific to mitochondrial proteins. Monitoring delivery of mitochondria to the lysosome Autophagosomes mature into autolysosomes upon fusion with the lysosome and for this reason, studies of mitophagy often employ lysosomal markers such as FPs targeted to the lysosome,35 antibodies against resident proteins of the lysosome such as catalase35 or lysosome-associated membrane protein 1 (LAMP1)36 to visualize the delivery of mitochondria to the lysosome. Lysosomes can also be labeled with the lysosomotropic LysoTracker® dyes.21, 29,37 Figure 3 demonstrates the use of a lysosomal marker, LAMP1-GFP, with MitoTracker® Red to visualize the co-localization of mitochondria within lysosomes in A549 cells. In this example, cells were 11 treated with 20 µM CCCP for 6 hours prior to confocal imaging. Studies of this nature often use a line profile analysis23,38 as demonstrated in Figure 3B. This approach allows for a correlation of intensity peaks that mirrors co-localization of fluorescent molecules. In this example, it is possible to see the dual LAMP1-GFP (green) peaks representing the lysosomal membrane and the single MitoTracker® Red peak illustrating the presence of a mitochondrion in a lysosome. In another model system, yeast, targeted GFP and RFPs27, 39, 40 as well as the FM family of dyes26,3941 or CMAC-Arg42 were used to label the vacuole (the yeast equivalent of the lysosome) and show delivery of labeled mitochondria to this structure. The relationship between lysosomes and autophagosomes is crucial to complete autophagic flux. This relationship breaks down if the lysosome fails to acidify (this impairs the functioning of lysosomal hydrolases and blocks autophagy). Failure of lysosomal acidification has been demonstrated to contribute to forms of inherited Alzheimer disease.43 Alternatively if fusion between the lysosome and the autophagosome is inhibited, incomplete autophagy would occur. While this scenario has not been extensively studied and is therefore poorly understood, dysfunction in autophagosomal-lysosome fusion may prove to be a critical aspect of autophagy in a number of diseases. For this reason, probes that allow simultaneous visualization of autophagosomes, autolysosomes and lysosomes within the same preparation are of huge benefit to researchers. Multi-color imaging aids in the discrimination of deficits in sequestration of (damaged) mitochondria to the autophagosome, a deficit presumably specific to mitophagy, versus deficits in fusion and/or acidification of the lysosome which would affect autophagy in general. 12 With this in mind, we recently developed a novel, red shifted lysosomotropic dye, LysoTracker® Deep Red (Zhou WJ, Dolman NJ, Smith RK, Gee K and Janes MS, unpublished data) that enables multi-color imaging of the accumulation of autophagy-specific markers in the lysosome (Dolman NJ, Chambers KM and Janes MJ, unpublished data). This novel probe can also be used to dissect aspects of mitophagy by adding an additional fluorescence detection channel to a given experimental protocol. Lysosomes can be detected in the ‘Deep Red’ channel (excitation and emission specifications approximately that of Alexa Fluor® 647 or Cy5 dyes) leaving the green (GFP/FITC) and red (TRITC/RFP) open for mitochondrial probes and markers of the autophagic machinery. Figure 4 shows three color imaging of mitophagy where mitochondria (Mito-RFP), SQSTM1 (GFP-SQSTM1) and lysosomes (LysoTracker® Deep Red dye) have been labeled under conditions favoring mitophagy. Under these conditions, mitochondria that are positive for SQSTM1 can be seen to co-localize with lysosomes. FP biosensors that utilize the pH difference between autophagosomes and autolysosomes have recently been developed to discriminate between these two structures. These constructs encode both a green and a red FP fused to the N-terminus of MAP1LC3B. These probes exploit the greater pH sensitivity of Aequorea victoria-derived GFP versus the red FPs, typically derived from Anthozoas (Discosoma striata or Entacmaea quadricolor. GFP (from Aequorea victoria) has a pKa of 6.1544 and is the most commonly used GFP. Anthozoa FPs have a pKa around 4.545 and therefore fluorescence from RFPs will typically be maintained despite the acidic environment of the autolysosome while GFP will lose fluorescence in this pH range. Tandem MAP1LC3 chimeras exploiting this differential acid-sensitivity have been constructed using GFP 13 and either mRFP46, tagRFP (Dolman NJ, Chambers K, Batchelor RH and Janes MS, unpublished data) or mCherry.36 The use of tandem fluorescent protein sensors is complicated by fluorescence emission overlap between the two FPs and therefore, researchers should be careful to avoid bleed-through of signal from one fluorescent protein into the detection channel of the other47 as well as the possibility of intra-molecular fluorescence resonance energy transfer (FRET) between the two FPs.48 The progression toward autolysosomal formation can also be monitored using lysosomal probes such as the LysoTracker® dyes and dextrans conjugated to fluorescent dyes . Dextrans are efficiently trafficked to lysosomes in live cells.49 If these dextrans are conjugated to spectrally resolvable fluorescent molecules such as Cascade Blue®46 or Alexa Fluor® 64736, the tandem FP MAP1LC3 chimera may be used to simultaneously image autophagosomes, autolysosomes and lysosomes. A similar approach using near infrared emitting mitochondrial dyes such as MitoTracker® Deep Red FM is appealing in that this combination would enable multi-color imaging of the sequestration of mitochondria into an autophagosome and subsequent maturation of these mitochondria-containing vesicles into autolysosomes. Measuring mitochondrial membrane potential As the loss of mitochondrial membrane potential is a trigger for mitophagy, potentiometric probes are commonly used to study mitophagy. A number of dyes including rhodamine 123, TMRM, TMRE, DiOC6 and JC-1 have been used to measure mitochondrial membrane potential.50 Of these, TMRM31,51, 52 or TMRE53-55 are the most commonly used dyes. As neither TMRE nor TMRM are well-retained in mitochondria following depolarization their utility in 14 longer term applications is limited. There are important considerations to be made when using TMRM or TMRE which are beyond the scope of this review. For a full discussion of these considerations, see Duchen et al.50 The dye JC-1 (5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide) offers a ratiometric means to measure mitochondrial polarization and has been used to study mitophagy.56-58 However, care must be taken to avoid spectral overlap between the two species and also to allow for equilibration of the dye across the plasma and mitochondrial membranes during cell loading. The phenomenon of J aggregate formation is also highly concentrationdependent and therefore optimization of the loading protocol is required. Image analysis and quantification of mitophagy As outlined above, many studies in the field of mitophagy involve a determination of colocalization between mitochondrial labels and markers for autophagy (for example MAP1LC319, 21, 23, 29, 30 ) or other salient proteins such as PARK230 ,31 or PINK130 The recruitment of mitochondria has also been analyzed through co-localization of other autophagy markers such as ATG8 or GABARAPL1 with Atg3226 ,27 or BNIP3L,15 respectively. Quantification of the extent to which markers co-localize provides a numerical index that can be subjected to statistical analysis. By employing this approach, studies have quantified the co-localization of MAP1LC316, 19, 29 or Atg827 on mitochondria. Alternatively, signaling molecules involved in mitophagy, such as PINK154 and PARK238, 52, 53 have been quantified according to the degree to which they co-localize with mitochondrial probes. Co-localization can be determined through pixel-by-pixel analysis of an image or from a line profile drawn across a cell/intracellular 15 structure. Both methods can be assessed for statistical significance using a suitable statistical test (for example the Pearson product-moment correlation coefficient54). There are a number of important considerations when interpreting co-localization. These include fluorophore/filter properties and acquisition mode. Filters with poor properties or fluorophores with broad excitation/emission properties may lead to fluorescence emission overlap where the emission from one fluorophore appears in both detection channels leading to false colocalization. This can be avoided through selection of fluorophores with suitable optical properties as well as by exciting each fluorophore separately or determining the extent of fluorescence emission overlap with samples labeled with one fluorophore only. Many modern laser-scanning confocal systems have acquisition configurations that allow the excitation of, and the emission from, each fluorophore to be collected sequentially. Wide-field microscopes employing excitation and emission filters mounted on a filter wheel dictate that the excitation of each fluorophores and subsequent emission collection occurs independently for each fluorophore. While this approach avoids false positives, there is the possibility of falsely negative co-localization when imaging motile structures in live-cells especially if the acquisition rate is too slow. Two structures that co-localize may move between each acquisition resulting in falsely negative co-localization. In addition to co-localization, studies on mitophagy also commonly quantify morphological and/or functional changes associated with the autophagic clearance of these organelles. Changes in mitochondrial morphology involve scoring cells based upon the appearance of mitochondria. The changes may be relatively simple, such as mean length,59 to more advanced measurements 16 of morphology based on mitochondrial phenotypes19, 60 or mitochondrial aggregation through a compaction index.61 Mitochondrial degradation related to the induction of mitophagy has been shown through the use of MitoTracker® Green54 as well as antibodies against the mitochondrial resident protein TOMM20/TOM20.38, 54, 62 Future directions Dissecting the early events in mitophagy The majority of studies described above relied upon co-localization of mitochondrial fluorescent probes with MAP1LC3. It should be noted that MAP1LC3 is not the only marker for autophagy and other proteins that participate in the autophagic machinery may also serve as useful markers for autophagy. A number of these alternate markers are amenable to fluorescence imaging (for a full review see ref. 28 and references in therein) and are particularly attractive as they provide delineation of early points in the autophagic pathway. These include ATG5-ATG12, ATG14, unc-51-like kinase 1(ULK1), ATG16L1, WD repeat domain, phosphoinositide interacting (WIPI), and zinc finger, FYVE domain containing 1 (ZFYVE1)/ Double FYVE-containing protein 1 (DFCP1).63 The order in which these proteins are recruited to the autophagosome has been characterized 63 and therefore it may be possible to observe damaged mitochondria being encompassed by structures positive for any of these markers which would serve to define how early in the process mitochondria, destined for mitophagy, become associated with the autophagosome. While there is a paucity of data on this area, early time points in the mitophagy pathway would be predicted to show co-localization with many of these markers and these studies would provide useful 17 information on the early stages of mitochondrial targeting and sequestration by the forming autophagosome. Furthermore, co-localization of any of these markers with mitochondrial probes and probes for key pro-mitophagy signaling proteins such as PINK1, PARK2, BNIP3, BNIP3L or SQSTM1 would further enable the elucidation of events that occur during mitophagy. FP-based biosensors for mitophagy The field of FP engineering is accelerating with a plethora of new variants described each year. These proteins show differential properties, many of which may provide useful tools to study mitophagy. For example, FP-based biosensors such as a pH-sensitive mitochondrial-targeted FP chimera has been recently developed to monitor mitochondrial accumulation in the yeast vacuole.42,64 This approach utilized the pH-sensitive GFP variant pHlourin65 in a tandem fusion with the pH-insensitive DsRed which has an acidic pKa.45 When degraded, yeast mitochondria localize to the acidic vacuole and the emission from pHluorin is lost leaving only the DsRed fluorescence emission and an indication of mitophagy. Outside the vacuole however, the mitochondrial-targeted tandem-FP biosensor is in a neutral pH environment and therefore fluorescence emission is derived from both DsRed and pHluorin. This new probe represents a particularly useful tool to study mitophagy however employing the dual FP (DsRed and pHluorin) approach in Rosella may have the caveats associated with other tandem FP biosensors, where the FPs may undergo FRET,48 exhibit fluorescence emission overlap or differential bleaching kinetics and brightness.47 An alternative strategy that circumvents many of the potential problems associated with tandem FP biosensors is to use a single FP-based biosensor with spectral properties that shift with changes in pH. The Miyawaki laboratory48 used the FP Keima to construct a mitophagy probe. Keima exhibits a pH dependent 18 shift in its peak excitation wavelength. At a high pH Keima absorbs light at 440nm whereas at acidic pH Keima absorbs light at a peak wavelength of 586 nm. Keima was successfully targeted to mitochondria48 yielding a FP-based biosensor for mitophagy. Under basal conditions the mitochondrial targeted Keima is in a neutral environment and absorbs 440 nm light. Upon induction of mitophagy, the probe accompanies mitochondria sequestered into autophagosomes and on to the lysosome. In the acidic lysosomal environment Keima becomes ionized and the peak absorption shifts to 586 nm. The authors show eloquent, quantitative imaging of mitophagy by measuring the ratio of emission intensities at 620 nm from the two excitation peaks. Furthermore, as this probe exhibits a large Stokes shift (the difference in wavelength between the excitation and emission wavelength), the authors note that Keima is ideally suited to multi-color imaging and demonstrate this through the three color imaging of mitochondrial-targeted Keima and an Alexa Fluor® 488 labeled dextran.48 Fluorescent probes to image individual mitochondria Recently, a subfamily of FPs known as ‘optical highlighters’ has been developed that allow precise visualization of a given subcellular structure. These optical highlighters comprise targeted FPs that can either be activated from a dark state to a fluorescent state,66 reversibly switched on and off,67 or converted from a fluorescent species with excitation and emission properties that differ from the post-irradiated fluorophore.68 Since these original reports, additional variants of these optical highlighters have been developed and are described in a comprehensive review of recent advances in fluorescent protein applications.69 These various optical highlighters, in particular photoactivatable GFP, have been used to study mitochondrial dynamics and have proven invaluable to study mitochondrial connectivity,60, 70, 71 as well as mitochondrial fission16 and mitochondrial elongation.72 These probes have helped describe the 19 role of mitochondrial dynamics in regulating mitophagy.16,72 The possibility afforded by these probes to label an individual mitochondrion and follow it through its life to its degradation in the lysosome is an attractive one in terms of monitoring mitophagy. Novel fluorescent probes Improvements in the spectral properties of fluorescent proteins to shift their fluorescence emission into the near infrared and infrared spectrum73, 74 may facilitate in vivo imaging of mitophagy. In vivo studies have already been performed using GFP-MAP1LC375 and combining these tools with mitochondrial-targeted FPs will aid our understanding of the dynamic processes of mitophagy that occur in whole organisms. An improved palette of FPs also aids in multi-color imaging where the simultaneous monitoring of three or four different parameters can be made using blue, green, red and near infrared fluorophores. This is already possible using near infrared-emitting dyes.21, 36 New probes such as LysoTracker® Deep Red (figure 4) will aid in the imaging of the delivery of multiple entities to the autolysosome. The development of additional longer wavelength probes is particularly attractive given the potential for multiparametric analysis of mitophagy it could enable. Imaging mitophagy with improved spatial resolution Fluorescence imaging has, until recently, been limited in resolution to the diffraction limit of light, around 250 nm.76 This has restricted the spatial detail that can be derived from a fluorescence image. Several parallel approaches have produced technologies that have broken the diffraction limit of fluorescence imaging; these are known as sub-diffraction limited imaging or super resolution microscopy (For a review see ref. 76). With these improved imaging 20 modalities fluorescence microscopy can achieve a resolution of 10 nm. 76 The use of these high resolution imaging modalities will provide further information concerning changes in mitochondrial structure as well as enhanced profiling of the co-localization and dynamics of mitochondria with elements of the autophagic machinery. Using light to trigger mitophagy Many experimental manipulations that trigger mitophagy often involve the loss of mitochondrial membrane potential, particularly when protonophores such as CCCP are used. However the use of CCCP causes a global loss of mitochondrial membrane potential and therefore has potential off-target effects.23 For this reason, the targeted depolarization or damage of a limited number of mitochondria presents a more physiologically relevant mechanism by which to stimulate mitophagy.23 The discovery and cloning of light-activated ion channels, known as channel rhodopsins, has led to advances in the study of electrically excitable cells given the ability to precisely and non-invasively depolarize a subset of cells in a field.77 It may be possible to target channel rhodopsin to subcellular locations such as mitochondria to aid in addressing mechanistic questions related to mitophagy.78 The specific expression of these channels in mitochondria offers an attractive option of controlling mitochondrial function78 and the clearance of mitochondria during autophagy. Certain fluorophores are particularly prone to produce reactive oxygen species upon photoactivation. Exploitation of this photosensitizing property represents a potentially useful approach to induce mitophagy through targeted damage. These highly reactive species rapidly react with proteins surrounding the photo sensitizer and the damage by ROS is spatially confined to the immediate vicinity of the sensitizer. This technique is known as chromophore-assisted light 21 inactivation (CALI).79 CALI can be used to specifically damage proteins in a spatially localized region80 through targeted ROS production.81 The use of CALI has been restricted by the difficulty of targeting the photosensitizing dyes to discrete cellular sites. Recently, Killer Red, a fluorescent protein that produces a burst of ROS upon illumination with 585 nm light has been described.82 KillerRed targeted to mitochondria was recently used to generate ROS in the mitochondrial matrix and study the role of oxidative stress and damage in the induction of mitophagy.23 Increased throughput for mitophagy assays Finally given the role of defective mitophagy in disease13,14 probes that accelerate the screening of compounds that modulate mitophagy constitute attractive therapeutic agents.83 To date, autophagy-specific assays that are amenable to high-throughput screening employ time-resolved FRET (TR-FRET) between GFP-MAP1LC3B and an anti-MAP1LC3B antibody specific to the cleaved form of MAP1LC3 (MAP1LC3-II). Using this approach, it is possible to detect either the activation or inhibition of autophagy.84 Currently there are no means to transfer this technique to study mitophagy but this is clearly an area that would benefit from further work. 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Maintenance of a healthy pool of mitochondria affects overall cell/organism viability. A change in mitochondrial health form a healthy pool of functional mitochondria (‘green’) to a pool containing a mitochondrion with impaired function (‘red’) can have a profound effect on the overall health of a cell or even an organism. Mitophagy functions as a protective mechanism whereby damaged mitochondria are encompassed by the nascent autophagosomes for subsequent breakdown in the autophagolysosome. This targeted degradation maintains a healthy pool of mitochondria and a complete pool of mitochondria is re established through mitochondrial fission. If this process is impaired or overwhelmed, the extent of mitochondrial dysfunction may be perpetuated, resulting in a greater number of damaged mitochondria (‘grey’). These mitochondria overwhelm the degradative capacity of the mitophagic pathway and proceed to impair cellular function, possibly triggering cell death (Bottom right). Mitochondria gate the mechanism by which cells die: via apoptosis or necrosis and the initiation of one pathway versus another has a profound effect on the manner in which that dead cell is dealt with by phagocytes and the extent to which an inflammatory response is mounted or circumvented. Figure 2. Co-localization of mitochondria and the autophagosomal marker MAP1LC3B. A mitochondrial-targeted red fluorescent protein (AII, BII) and GFP-MAP1LC3B (AI, BI) were expressed in HeLa cells using BacMam gene delivery 48 hours after transduction, cells were treated with vehicle (A) or 10 µM CCCP/60 µM chloroquine (B, used to inhibit autophagic flux and cause the accumulation of autophagosomes) and cultured overnight. Depolarization of mitochondria with CCCP was strongly associated with co-localization of GFP-MAP1LC3B on RFP-labeled mitochondria (BIII) versus control conditions (AIII). The regions indicated in dashed white boxes in AIII and BIII are enlarged in AIV and BIV respectively. Co-localization 33 of GFP-MAP1LC3B and Mito-RFP is shown by the arrow heads in BIV. Cells were imaged on a Zeiss LSM 710 Confocal microscope using a water immersion 40x 1.4 NA lens and appropriate filter sets. Scale bar =10 µm (AI-III, BI-III) and 2 µm for AIV and BIV. Figure 3. Lysosomal degradation of mitochondria. A lysosomal-targeted GFP (LAMP1) was expressed in HeLa cells using BacMam gene delivery. Cells were then incubated with 10 µM CCCP in the presence of 30 µM chloroquine (to block autophagic flux) and loaded with 50 nM MitoTracker® Red in DPBS for 30 minutes at 37o C. Cells were imaged on a Zeiss 710 Laser Scanning Confocal microscope using a water immersion 40x 1.4 NA lens and appropriate filter sets. Under conditions favoring mitophagy, numerous mitochondria could be seen co-localized with lysosomes (AI). Enlargement of one of these structures (A II) shows the lysosome (AII) surrounding a mitochondrion (AII, AIV). A line profile (B) through this structure further highlights this encompassment and are powerful analytical tools to examine the presence of molecules within subcellular structures . Cells were left for 48 hours after transduction and before drug treatment and imaging. Scale bar =10 µm for AI and 5 µm for AII-III. Figure 4. Accumulation of SQSTM positive mitochondria in lysosomes following CCCP treatment. A mitochondrial targeted RFP (red) and GFP-SQSTM1 (green) were expressed in HeLa cells using BacMam gene delivery. The following day cells were loaded with 50 nM LysoTracker® Deep Red (blue) in complete media for 15 minutes at 37oC (A). Cells were imaged on a Zeiss 710 Laser Scanning Confocal microscope using a water immersion 40x 1.4 NA lens and appropriate filter sets. Cells were then treated with 10 µM CCCP for 6 hours to depolarize mitochondria. During the CCCP incubation period HeLa cells were also treated with 90 µM Chloroquine to block autophagic flux and thereby favoring mitochondria positive 34 lysosomes. The region indicated by the white dashed box in A is enlarged in BI-IV. Fragmented mitochondria (BII) are observed co-localizing with punctate GFP-SQSTM1 staining (BI, BIV) and Lysosomes (BIII & BIV). Scale bar =5 µm. 35