Autophagy is an evolutionarily conserved lysosome-mediated degradation and recycling process, which functions in cellular homeostasis and stress adaptation. The process is highly dynamic and involves autophagosome synthesis, cargo recognition and transport, autophagosome–lysosome fusion, and cargo degradation. The multistep nature of autophagy makes it challenging to quantify, and it is important to consider not only the number of autophagosomes within a cell but also the autophagic degradative activity. The rate at which cargos are recognized, segregated, and degraded through the autophagy pathway is defined as autophagic flux. In practice, methods to measure autophagic flux typically evaluate the lysosome-mediated cargo degradation step by leveraging known autophagy markers such as MAP1LC3B (microtubule-associated proteins 1A/1B light chain 3 beta) or lysosome-dependent fluorescent agents. In this review, we summarize the tools and methods used in mammalian cultured cells pertaining to these two approaches, and highlight innovations that have led to their evolution in recent years. We also discuss the potential limitations of these approaches and recommend using a combination of strategies and multiple different autophagy markers to reliably evaluate autophagic flux in mammalian cells.
Autophagy is a highly conserved eukaryotic degradation and recycling process whereby cytoplasmic material is degraded via the lysosome. The most well-studied form of autophagy, macroautophagy, utilizes double-membrane vesicles, autophagosomes, to deliver cytoplasmic contents to the lysosome for degradation, and is the subject of this review. Macroautophagy (hereafter referred to as autophagy) has widespread functions in normal development and in pathological conditions, including pathogen infections, diabetes, neurodegenerative disorders, and cancer [1–3]. These disease links have led to escalating interests in studying the autophagy process in a variety of contexts, with a corresponding need for robust strategies to accurately monitor autophagy. In particular, it is important to consider not only the presence or number of autophagosomes but also autophagic flux. The latter is defined as the completion of the autophagy process, including cargo recognition, synthesis of autophagosomes, transport of cargo-laden autophagosomes to lysosomes, fusion of autophagosomes and lysosomes, degradation of autophagic cargo, and release of degradation products .
During the autophagy process, membrane nucleation results in the formation of the cup-shaped phagophore, or isolation membrane, which subsequently elongates and sequesters cargos. Sealing of the elongated membrane forms a vesicular autophagosome that fuses with lysosomes to mediate cargo degradation (Figure 1). The core proteins required for the autophagy process are designated as autophagy-related (ATG) proteins. Two ubiquitin-like conjugation systems, ATG12 (autophagy-related 12)–ATG5 (autophagy-related 5)–ATG16L1 (autophagy-related 16 like 1) and MAP1LC3B (microtubule-associated proteins 1A/1B light chain 3 beta; LC3B)–ATG7 (autophagy-related 7)–ATG3 (autophagy-related 3), are required for membrane elongation. LC3B (a homolog of yeast Atg8) is first cleaved by the cysteine protease ATG4B (autophagy-related 4B cysteine peptidase) to expose the C-terminal glycine and form LC3B-I, which is then transferred by ATG7 and ATG3, and finally conjugated by the ATG12–ATG5–ATG16 complex to phosphatidylethanolamine (PE) on the elongating membrane. Membrane-bound LC3B is termed LC3B-II. The membrane continues to expand and curve, sequestering cytoplasmic cargos inside followed by sealing of the membrane ends to form the double-membrane autophagosome . Cleavage and recycling of LC3B-II from the outer membrane, again by ATG4B, is a step that was shown to be required for lysosomal fusion in yeast , but such a requirement in mammals has not yet been determined. In mammalian cells, the N-ethylmaleimide-sensitive factor attachment protein receptor (SNARE) Syntaxin 17 (STX17) localizes to the outer membrane of the fully sealed autophagosome and promotes autophagosome–lysosome fusion [7,8]. Autophagosome fusion with lysosomes results in the formation of autolysosomes . Following the fusion, the acidification and degradation of the inner autophagosomal membrane leads to release of STX17 from its outer membrane . Within the autolysosome, the acidic hydrolases degrade the cargos into small molecules, such as amino acids, lipids and carbohydrates, which are returned to the cytoplasm by lysosomal permeases [9–11] (Figure 1).
An overview of the autophagy process and the life cycle of MAP1LC3B (LC3B).
While all the above steps are important for the autophagy process and can contribute to the levels of autophagic flux, most methods to monitor flux rely primarily on evaluating autophagosome–lysosome fusion and/or autophagic cargo degradation. The reason for evaluation of these steps is to distinguish between autophagosome numbers and degradative activity, and to determine whether the autophagy process has reached the degradative step. For the purpose of this review, we use the term autophagic flux to refer primarily to autophagic degradation or autophagosome–lysosome fusion, with the latter assuming that autophagic cargos would be degraded by lysosomal hydrolases. Autophagy markers, such as LC3B and lysosome-dependent fluorescent agents, are two common routes by which autophagic flux is assessed. We summarize here tools and methods, with particular focuses on the roles of autophagy markers and lysosome-dependent fluorescent agents, commonly used to monitor autophagic flux in mammalian cultured cells, and highlight recent developments and new techniques that have improved upon the established strategies.
Monitoring autophagic flux through autophagy markers
Several of the core autophagy proteins serve as useful indicators or markers for the autophagy process itself. These autophagy markers localize to autophagosomes such that they can be used to label the double-membrane vesicles and/or are degraded alongside autophagic cargos, and thus can be quantitated through immunoassays to infer autophagic flux. One of the most widely monitored proteins is the aforementioned LC3B. Lipidated LC3B-II lines both the inner and outer membranes of autophagosomes. During the late stage of the autophagy process, the outer LC3B-II is removed by ATG4B, whereas the inner LC3B-II is degraded by lysosomal proteases (Figure 1) . These aspects of LC3B make it an excellent marker for monitoring autophagic flux and have spurred the development of numerous tools and methods currently used to evaluate autophagic flux [13,14].
Other autophagy markers include a wide variety of autophagy receptors (ARs). Traditionally, autophagy was considered as a bulk degradative pathway with no selectivity toward substrates, but more recent studies have discovered an array of substrates that are preferentially degraded . These substrates are targeted using ARs, which physically link, through their LC3-interacting region (LIR)-motif, the substrates to LC3B. ARs, such as p62/SQSTM1 (sequestosome-1) and NBR1 (neighbor of BRCA1), recognize substrates in an ubiquitin-dependent manner (Figure 1) . Like LC3B, the ARs themselves are autophagy substrates and are degraded in autolysosomes . Several types of selective autophagy have been discovered and include the degradation of specific organelles such as mitochondria (mitophagy), endoplasmic reticulum (ER-phagy or reticulophagy), and the nucleus (nucleophagy), as well as the degradation of foreign objects such as bacteria (xenophagy) and viruses (virophagy) [18–21]. A comprehensive but not exhaustive list of the ARs that have been identified for different types of selective autophagy is summarized in Table 1.
|Type of autophagy||Selective substrates||Autophagy receptor||Reference(s)|
|DNA-mediated xenophagy||Mycobacterium tuberculosis||p62|||
|(Endoplasmic) reticulophagy||Endoplasmic reticulum||RETREG1 (FAM134B)|||
|Midbody ring disposal||Midbody protein CEP55||p62||[61,62]|
|Virophagy||HIV p24 and viral proteins||TRIM5α||[69,70]|
|Xenophagy||Bacteria and foreign objects||Galectin 8||[71–73]|
|Type of autophagy||Selective substrates||Autophagy receptor||Reference(s)|
|DNA-mediated xenophagy||Mycobacterium tuberculosis||p62|||
|(Endoplasmic) reticulophagy||Endoplasmic reticulum||RETREG1 (FAM134B)|||
|Midbody ring disposal||Midbody protein CEP55||p62||[61,62]|
|Virophagy||HIV p24 and viral proteins||TRIM5α||[69,70]|
|Xenophagy||Bacteria and foreign objects||Galectin 8||[71–73]|
Abbreviations: BCL2L13, BCL-2-like 13; BNIP, BCL-2/adenovirus E1B 19 kDa interacting protein 3; CEP55, centrosomal protein of 55 kDa; FUNDC, FUN domain-containing protein; HDAC6, histone deacetylase 6; NBR, next to BRCA gene 1; NCOA4, nuclear receptor co-activator 4; NDP52, nuclear dot protein 52; NIX, NIP3-like protein X; OPTN, optineurin; SMURF, SMAD ubiquitylation regulatory factor; STBD1, starch-binding domain-containing protein 1; TAX1BP, Tax1-binding protein; TECPR1, tectonin β-propeller repeat-containing protein; TOLLIP, Toll-interacting protein; TRIM, tripartite motif.
A relatively new autophagy marker is STX17, which localizes to sealed autophagosomes, but not open isolation membranes, and functions in autophagosome–lysosome fusion . Using live cell imaging, green fluorescent protein (GFP)-STX17-labeled structures were shown to fuse with the lysosome marker mCherry-LAMP1 . Using a cyan fluorescent protein (CFP)–STX17–TM (transmembrane) construct and time-lapsed imaging, Tsuboyama et al.  showed that STX17 localized to sealed autophagosomes and was then released following autophagosome–lysosome fusion, inner autophagosomal membrane degradation, and autolysosome acidification. The analysis of STX17 dynamics in conjunction with lysosome markers is a promising new flux assay, bearing in mind that STX17 also localizes to the ER and mitochondria  and that alternate SNAREs may be utilized in some autophagy contexts [24,25].
Immunoassays for assessing turnover of autophagy markers
One common approach to evaluate autophagic flux in mammalian cells is to measure the turnover of autophagy markers. As described above, these markers are also substrates of the autophagy process, and measuring their turnover using immunoblotting is a simple and inexpensive method to determine the relative level of autophagic flux. This method has been extensively reviewed [13,14] and so is not discussed in great detail here. Briefly, a decrease in AR levels by immunoblot analysis suggests an increase in autophagic flux and vice versa. Assessing autophagic flux using LC3B as a marker is somewhat distinct due to its dynamic involvement in the autophagy process and typically involves measuring LC3B-II level in the presence or absence of a lysosomal inhibitor to determine whether autophagy flux is enhanced, reduced, or blocked. A further increase in LC3B-II level in the presence of the lysosomal inhibitor would suggest that the given context or treatment increases autophagic flux, whereas failure to further increase LC3B-II level in the presence of the lysosomal inhibitor would suggest that the context or treatment in question inhibits autophagic flux .
In addition to immunoblot detection of LC3B-II turnover, LC3B-enzyme-linked immunosorbent assay (LC3B-ELISA) following subcellular fractionation was recently employed to assess autophagy levels through the differential subcellular localization (i.e. membrane-bound, mitochondrial, or cytosolic) of LC3B proteins. The ratio of LC3B proteins in membrane versus cytosolic fractions was used to reflect the proportion of LC3B engaged in the autophagy process and can be used to measure relative autophagy levels across different conditions in cells or tissues. Like the LC3B-II immunoblot turnover assay, LC3B-ELISA allows the relative measurement of autophagic flux by detecting LC3B-II level in the presence or absence of lysosomal inhibition. Through direct comparison of LC3B-ELISA and LC3B-II immunoblot, the authors showed that the LC3B-ELISA assay has advantages over the immunoblot assay in that it requires less antibodies, and has greater sensitivity, reproducibility, and multiplexing capability . The authors also showed that detection of mitochondrial-to-cytosolic LC3B levels following mitophagy induction yielded results comparable to using immunoblot. The capability of this method in detecting mitophagic flux suggests it may also be applicable to the study of other selective autophagy processes that target damaged organelles. The multiplexing capability of the ELISA platform supports simultaneous quantitation of additional protein markers such as other mammalian homologs of yeast Atg8 [see below regarding MAP1LC3 (microtubule-associated proteins 1A/1B light chain 3) and GABARAP (GABA type A receptor-associated protein) subfamilies] as well as ARs in different subcellular fractions. This aspect can be leveraged to detect and distinguish the activation of selective autophagy process(es) of interest. For example, FAM134B (RETREG1; reticulophagy regulator 1) is an AR of ER-phagy, while BNIP3 (BCL2-interacting protein 3) and NIX (BNIP3L; BCL2-interacting protein 3 like) are ARs of mitophagy (Table 1), and quantitating these ARs in addition to MAP1LC3s and GABARAPs across ER, mitochondrial, and cytoplasmic fractions could distinguish activation of ER-phagy versus mitophagy. This method, however, requires sufficient knowledge of protein markers' specificity, reliable and specific antibodies, and successful isolation of subcellular fractions of interest.
Important considerations for autophagy marker turnover assays
Many of the discovered ARs have autophagy-independent functions and can be degraded by other pathways such as the ubiquitin–proteasome system; therefore, their turnover should be interpreted with caution. For example, p62 is involved in many activities including cell death, oxidative stress, inflammatory response, and proteasomal degradation, which could affect p62 level in an autophagy-independent manner . Evaluation of autophagic flux based solely on changes in the level of a single AR could therefore be misleading. Lysosomal inhibitors such as Bafilomycin A1 (BAF) and chloroquine (CQ) are frequently used to discern the turnover of ARs by autophagy from other degradation pathways, since autophagic degradation of the ARs should be blocked when lysosomes are dysfunctional. In addition, detection of multiple ARs and/or the inclusion of other autophagy markers such as LC3B in the turnover assay is recommended.
LC3B is, by far, the most studied and utilized autophagy marker, yet seven homologs of LC3B have been identified in mammals. Together, they form the MAP1LC3 and GABARAP subfamilies. The former is made up of MAP1LC3A, MAP1LC3B, MAP1LC3B2 (microtubule-associated proteins 1A/1B light chain 3 beta 2), and MAP1LC3C (microtubule-associated proteins 1A/1B light chain 3 gamma), while the latter family is made up of GABARAP, GABARAPL1 (GABA type A receptor-associated protein like 1), and GABARAPL2 (GABA type A receptor-associated protein like 2). A previous study showed that both of these subfamilies are required for autophagic flux, but the MAP1LC3 subfamily is responsible for elongation of the isolation membrane while the GABARAP subfamily is required for its closure . A more recent study employing knockout of six LC3/GABARAP family members in HeLa cells showed that this family is essential for autophagosome–lysosome fusion but, unexpectedly, is not absolutely required for autophagosome formation . Indeed, autophagosome-like structures were still generated in cells with defective ubiquitin-like conjugation systems, but these cells had significantly reduced autophagic degradative activity . In addition, both MAP1LC3 and GABARAP subfamily members were initially discovered outside the autophagy pathway and have non-autophagy functions . Hence, whether we can confidently determine changes in autophagy flux based on LC3B turnover alone is important to consider. For example, if certain contexts or autophagy conditions rely predominantly on other family members such as GABARAP, then evaluation of LC3B alone may lead to misinterpretation of overall effects on autophagic flux. For these reasons, we recommend employing multiple flux assays incorporating different autophagy markers. We also suggest using more precise terminology to report findings from any particular assays; e.g. specifying ‘LC3B-mediated autophagic flux’ rather than simply ‘autophagic flux’.
Another emerging scenario that requires attention is the interpretation of the LC3B turnover assay when studying ATG4B inhibition. With the rising interest in autophagy modulation, several studies have investigated the potential of ATG4B as a target for autophagy inhibition . As mentioned above, it is still unclear whether recycling of LC3B-II from the outer autophagosomal membrane is required for the fusion between autophagosomes and lysosomes in mammalian cells. An increase in LC3B-II in ATG4B knockdown conditions could therefore indicate a block in recycling of LC3B-II on the outer autophagosome membrane, but not necessarily indicate reduced autophagosome–lysosome fusion and turnover. Similarly, in ATG4B knockdown conditions, a further increase in LC3B-II level by a lysosomal inhibitor may not necessarily indicate enhanced autophagy but rather a block in ATG4B-mediated recycling of LC3B-II.
MAP1LC3/GABARAP-based fluorescent reporter assays
The preferential localization of LC3B to autophagic vesicles is leveraged to monitor autophagic vesicles and autophagic flux in mammalian cells. Different fluorescent protein-tagged LC3B constructs have been developed and are currently in use as reporters for monitoring the autophagy process. Fluorescently tagged LC3B follows the same path as endogenous LC3B, where it is cleaved by ATG4B, conjugated to PE on elongating autophagic membranes, and then incorporated into autolysosomes. The decoration of autophagic vesicles by these fluorescent LC3B proteins is detectable as punctate structures that can be visualized and quantitated via fluorescence microscopy [31,32]. Some of the established fluorescent protein-tagged LC3B constructs include GFP–LC3B, mRFP–GFP–LC3B (tfLC3B; tandem fluorescent-tagged LC3B), mCherry-EGFP-LC3B, and pHluorin-mKate2-human LC3 (PK-hLC3) that have been reviewed extensively in ref. . Briefly, GFP–LC3B consists of a GFP-tagged LC3B and was one of the earliest constructs developed for monitoring autophagic vesicles . The tfLC3B was derived from GFP–LC3B and contains an additional monomeric red fluorescent protein (mRFP). GFP is prone to fluorescent quenching and denaturation, whereas mRFP is considerably more stable in low pH environments such as in autolysosomes. Thus, tfLC3B can be used to distinguish autolysosomes from autophagosomes by the absence of green fluorescence. In essence, autolysosomes would be decorated exclusively with red fluorescence, whereas colocalization of red and green fluorescence would mark autophagosomes (Figure 2A) . Weak fluorescence of GFP under mildly acidic conditions led to the development of several modified tandem fluorescent-tagged LC3 reporters (reviewed in ref. ). For example, PK-hLC3 is an improved version of tfLC3B where pHluorin, with a higher pKA (∼7.6) than GFP (∼5.9), was employed and is better quenched in autolysosomes to allow for more accurate delineation between autophagosomes and autolysosomes . In addition to fluorescence microscopy, flow cytometry may also be utilized to infer autophagic activity based on fluorescence emitted from the fluorescent protein(s) of choice. For example, a previous study used tfLC3B with flow cytometry to infer autophagic flux by evaluating the percent of RFP-positive cells that were also GFP-positive . Because GFP is quenched and denatured in autolysosomes, a reduction in green fluorescence among RFP-positive cells would suggest an increase in autophagic activity and vice versa. However, contrary to fluorescence microscopy, which is capable of visualizing individual autophagic vesicles, flow cytometry enables high-throughput assessment of autophagy level in thousands to millions of cells in exchange for the resolution required to monitor individual autophagic vesicles. The optimal method for assessing autophagic flux using fluorescent protein-tagged LC3s ultimately depends on the desired resolution and throughput of the experiment on hand.
Fluorescent protein-tagged LC3B molecules serve as fluorescent reporters to monitor autophagosomes and measure autophagic flux.
It is important to be aware that the above reporter constructs do not provide a clear indication of whether a reduction in the fluorescent signals is a result of autophagic degradation or reduced expression of the reporter protein. This problem could be addressed with a more recently developed reporter construct, GFP–LC3B–RFP–LC3BΔG (LC3B with C-terminal glycine residue deleted). This construct consists of GFP–LC3B fused to the N-terminus of an RFP-LC3BΔG mutant. ATG4B cleaves the C-terminus of LC3B to expose a C-terminal glycine required for conjugation to PE on the autophagosomal membrane, and the deletion of this residue on RFP-LC3BΔG prevents it from being conjugated to PE and instead retains it in the cytosol. RFP-LC3BΔG is thus not incorporated into autophagic vesicles and can act as an internal control that reflects the overall expression of the GFP–LC3B–RFP–LC3BΔG construct in a cell. When GFP–LC3B–RFP–LC3BΔG is expressed in cells, ATG4B cleaves the C-terminus of the wild-type LC3B forming GFP–LC3B-I and releasing RFP–LC3BΔG to the cytosol. While RFP–LC3BΔG remains intact in the cytosol, GFP–LC3B-I can be conjugated to PE on autophagic vesicle membranes and a fraction of it degraded in autolysosomes (Figure 2B). Autophagic flux can therefore be measured by computing GFP/RFP signal ratios, either through fluorescence microscopy or flow cytometry. High GFP/RFP ratios indicate low autophagic flux, whereas low GFP/RFP ratios indicate high autophagic flux .
Use of transgenic constructs (e.g. tfLC3B) in cells for the tracking of autophagosomes or measurement of autophagic flux inevitably introduces extraneous proteins into the cellular repertoire. The presence of these exogenous proteins can result in unwanted biological consequences. For example, overexpressing LC3B can increase autophagic flux and can lead to autophagy-independent aggregation of LC3B . In addition, the fluorescent protein conjugated to the protein marker of interest may also affect the function of the fusion protein. A recent study found that GFP-tagged LC3s and GFP-tagged GABARAPs were functionally impaired during mitophagy and did not robustly translocate to mitochondria, but these impairments did not occur when GABARAPs were instead fused to the smaller hemagglutinin tag . To address these issues, two recently published studies exploited the LIR motifs of ARs to develop fluorescent sensors that detect endogenous LC3/GABARAPs. These sensors comprise a fluorophore along with a short peptide sequence either adapted from the LIR of a known AR or optimized from such a sequence. With an additional hydrophobic membrane recruitment or oligomerization domain, these fluorescent sensors preferentially bind to endogenous LC3-II and fluorescently label LC3-II-rich autophagosomes. Different MAP1LC3 and GABARAP family members localize and function differently throughout the autophagy process . In vitro characterization of cells expressing these fluorescent LC3/GABARAP sensors have found that they are able to preferentially detect different endogenous LC3/GABARAP family members depending on the LIR sequence incorporated, with minimal effects on autophagic flux [39,40]. This ability to preferentially bind to specific homolog(s) may be leveraged to track specific MAP1LC3/GABARAP proteins in cells. For example, mCh-AS3_67  and HyD-LIR(Fy)-GFP  are two sensors that preferentially localized to MAP1LC3C and MAP1LC3A/B-positive autophagosomes, respectively. These two sensors can be used concurrently to track the dynamics of autophagosomes decorated with MAP1LC3C versus MAP1LC3A/B in live cells. Detection of these sensors in combination with lysosomes using live cell imaging techniques may also open up new strategies for monitoring autophagic flux in real time. It is important to note that both studies failed to identify a specific LIR sensor that can distinguish LC3A from LC3B, or vice versa, indicating that some ARs may interact with both LC3A and LC3B. Stolz et al.  also noted that the negligible effects of overexpressing these LIR sensors on autophagic flux could be attributed to their weaker affinity to MAP1LC3/GABARAP proteins compared with the affinity between endogenous ARs and MAP1LC3/GABARAPs. Nonetheless, these LIR sensors are promising new tools that warrant further optimization and in-depth in vitro characterization to aid in tracking autophagosomes.
As with immunoassays for assessing autophagy markers to evaluate autophagic flux, lysosomal inhibitors such as BAF and CQ may be employed in conjunction with the above fluorescent reporters to ensure that the changes in fluorescence are due to changes in autophagic flux rather than those in fusion protein expression or aggregation. Using tfLC3B as an example, treatment with BAF or CQ would inhibit lysosomal acidity and activity, thereby preventing fluorescent quenching and denaturation of GFP, leading to an increase in GFP signals in addition to RFP signals. In other words, lysosomal inhibition should rescue the absence of GFP signal from tfLC3B if the turnover of tfLC3B is mediated through autophagy. Also, with the aforementioned caveats of LC3/GABARAP in monitoring the autophagy process, it is recommended to detect multiple LC3/GABARAP family members and/or ARs as well as utilizing different assays when measuring autophagic flux.
Monitoring autophagic flux through lysosome-dependent fluorescent agents
An alternative approach to assess autophagic flux is the use of fluorescent agents or fluorogenic substrates that selectively partition to autophagic vesicles, with lysosomes being the frequent target due to their distinct physical properties such as low pH. Depending on the desired resolution and throughput, fluorescent labeling by these dyes or substrates can be qualitatively and/or quantitatively detected through microscopy, spectrophotometry, or flow cytometry. Many of the fluorescent dyes have been recently reviewed in detail by Klionsky et al. . This section will first provide a brief overview of a few of the fluorescent agents commonly used for assessing autophagic flux: LysoTracker, DQ-BSA, and Cyto-ID followed by more detailed reviews of an emerging fluorescent protein called Keima and two novel fluorescent agents developed over the past year: HQO [2,6-bi(2-(3,3-dimethyl-1-propylindolin-2-ylidene)ethylidene)-cyclohexanone] and Lyso-OC [7-((4-Methoxyphenyl)ethynyl)-N-(2-morpholinoethyl)-2-oxo-2H-chromene-3-carboxamide].
LysoTracker, DQ-BSA, and Cyto-ID
Autophagosome–lysosome fusion results in the formation of autolysosomes with low internal pH. LysoTracker (ThermoFisher Scientific) is an acidotropic dye that accumulates and fluoresces in acidic compartments, and may be used to differentiate between autophagosomes and autolysosomes when used in combination with autophagosome-specific markers like LC3B . This allows for the evaluation of autophagic flux by performing ratiometric measurement with fluorescence microscopy analysis. DQ-BSA is a fluorogenic protease substrate-derived bovine serum albumin (BSA). DQ-BSA is endocytosed and digested by proteases in cellular compartments such as endosomes, lysosomes, and autolysosomes. Proteolysis of DQ-BSA results in the release of fluorescent BSA fragments that may be detected through fluorescence microscopy or flow cytometry, and used to infer the levels of autophagic flux but only when used, like LysoTracker, in combination with autophagosome-specific markers .
Cyto-ID (Enzo Biochem, Inc.) is a cationic amphiphilic fluorescent tracer dye developed from identification of titratable functional moieties that allows selective staining of autophagic vesicles but not lysosomes. Compared with acidotropic dyes or DQ-BSA, Cyto-ID allows specific labeling of vesicles involved in the autophagy process rather than generic staining of acidic compartments. The utilities of Cyto-ID in detection of autophagic vesicles and estimation of autophagic flux were meticulously examined by Guo et al.  and were shown to provide more sensitive and specific labeling of autophagic vesicles compared with acidotropic dyes. In addition, Cyto-ID is suitable for microscopic, cytometric, and spectrophotometric applications making it a compelling choice in experiments involving high-throughput screens . Nevertheless, while Cyto-ID selectively stains and allows the monitoring of autophagic vesicles, it does not provide direct measurement of autophagic flux, but can be used in combination with lysosomal inhibitors such as BAF and CQ to infer the relative level of autophagic flux between conditions or treatments.
Keima is an acid-stable, coral-derived RFP that is gaining popularity in autophagic flux assessment. Keima has a pH-dependent bimodal excitation spectrum with a large Stokes shift, making it a suitable tool for distinguishing autolysosomes from autophagosomes based on intravesicular pH and compatible with green-emitting fluorophores such as enhanced GFP (EGFP) [44,45]. When transfected into cells, Keima is resistant to lysosomal hydrolases and can be excited at 438 nm under neutral pH or 550 nm under low pH (<6) to emit red fluorescence that peaks at 610 nm. This differential excitation makes Keima a potential tool for tracking autophagosome–autolysosome conversion and deriving a cumulative quantitative readout of autophagic flux . To evaluate autophagic flux using Keima, conversion of autophagosomes into autolysosomes is measured through laser-scanning confocal microscopy with alternating lasers of 438 and 550 nm for dual-excitation ratiometric imaging. Acidic compartments would be marked with a high 550/438 nm ratio and presumably represent autolysosomes or lysosomes . Keima has certain advantages over LC3/GABARAP-based fluorescent reporters in that it does not depend on the presence of LC3/GABARAP on autophagic vesicles thereby avoiding caveats associated with LC3/GABARAP-based assays mentioned previously. Indeed, Keima was used to detect autophagy in Atg5−/− mouse embryonic fibroblasts, which have a defective autophagy ubiquitin-like conjugation system .
Like other fluorescent proteins, Keima can be fused to proteins or organelle-targeting peptides of interest to monitor the localization of specific proteins or organelles. Katayama et al.  generated mt-mKeima, a mitochondria-targeting Keima, by fusing Keima to a tandem repeat of cytochrome C oxidase III presequence. mt-Keima localizes to mitochondria and detects the conversion of mitochondria from neutral to low pH (i.e. mitophagy) through dual-excitation ratiometric imaging [44,46,47]. In theory, Keima can also be fused to any protein of interest to track the autophagy-mediated turnover of the target protein. However, it is unclear whether Keima specifically monitors autophagosome–autolysosome conversion, as microautophagy (another type of autophagy independent of autophagosomes) may directly transport Keima from the cytosol to lysosomes and lead to increased 550/438 nm . To more precisely determine whether a change to the 550/438 nm ratio of Keima is due to a change in (macro)autophagic flux, other markers such as LC3/GABARAP proteins or ARs may be tagged with a green fluorophore and visualized simultaneously with Keima.
Recent developments of fluorescent agents to assess autophagic flux have exploited the physical properties of lysosomes, specifically during successful fusion with autophagosomes, to precisely measure autophagic flux. An example of this is HQO, a cyanine dye developed for the evaluation of mitophagy flux. Mitophagy is commonly assessed through simultaneous fluorescent labeling of mitochondria and autophagosomes or lysosomes, where colocalization of both entities is indicative of mitophagy-autophagosome or lysosome-mediated mitochondria degradation, respectively. Such labeling can be achieved through protein markers and/or fluorescent dyes. This approach is often restricted by the limited selection of compatible fluorescent probes with minimal fluorescent emission spectral overlap and by the lack of protein markers specific for mitochondria and autophagosomes/lysosomes . HQO has excitation/emission (ex/em) wavelengths of 530/650 nm and specifically accumulates in mitochondria of live cells with minimal cytotoxicity (Figure 3A).
The mechanisms of two recently developed lysosome-dependent fluorescent agents: HQO and Lyso-OC.
Upon exposure to a reduced pH such as in the instance of delivery into autolysosomes, HQO is protonated to HQOH+, and its ex/em wavelengths redshift to 710/750 nm. This significant shift in the ex/em wavelengths allows distinction between mitochondria at physiological pH and those presumably in autolysosomes (Figure 3A). Ratiometric measurements of fluorescence emission from HQOH+/HQO can therefore be used to infer mitophagy flux . While HQO is specific for the assessment of mitophagy, its unique features of (1) localizing to a specific compartment (i.e. mitochondria) and (2) undergoing shift in ex/em wavelengths upon exposure to an acidic environment shed light on possible avenues for the development of dyes applicable for assessing autophagic flux in general. For instance, a chemical dye that combines the autophagic vesicle-localizing characteristic of Cyto-ID and the property of HQO in changing fluorescent ex/em wavelengths upon exposure to a lower pH would allow the monitoring of autophagosomes and evaluation of autophagic flux using a single chemical agent based on individual and ratiometric fluorescence measurements, respectively. Similar concepts may also be applied in developing dyes that specifically label other organelles to monitor their delivery to lysosomes for assessment of selective autophagic flux.
Another fluorescent agent recently developed leverages the change in lysosomal polarity upon autophagosome–lysosome fusion. Chemical polarity of a vesicle is dependent on the charge distribution of the constituents in the vesicular compartment. Because lysosomes and autophagosomes are different in their constituents (i.e. enriched with enzymes and cytosolic fractions, respectively), autolysosomes produced from their fusion have different chemical polarities from that of lysosomes, and this property can be utilized to discriminate autolysosomes from lysosomes . (Lyso-OC) is a two-photon dye comprised of polarity-dependent fluorescent coumarin and lysosome-targeting morpholine moieties. Together, these moieties allow Lyso-OC to selectively partition to lysosomes and, upon two-photon excitation at 760 nm, to emit fluorescence at 490–550 nm in lysosomes. The polarity-dependent fluorescent property of the coumarin moiety is quenched in autolysosomes, and this allows Lyso-OC to specifically stain lysosomes and not autolysosomes (Figure 3B). The capability of Lyso-OC in assessing autophagic flux in cells was supported by the reduction in fluorescence at 490–550 nm upon nutrient starvation. Treatment with 3-methyladenine (3-MA), an early-stage autophagy inhibitor, was able to abrogate the effect of nutrient starvation on Lyso-OC fluorescence . In addition, the probe was not cytotoxic, making it suitable for live cell imaging and potentially applicable to flow cytometry for high-throughput screens. Although Lyso-OC selectively and exclusively stains lysosomes and therefore cannot be utilized to specifically monitor autophagic vesicles, its capability in detecting changes in autophagic flux through diminished fluorescence renders it a compelling and alternative tool for evaluating autophagic flux in live cells and tissues. However, caution should be exercised when using this dye to measure autophagic flux, as other cellular processes or lysosomotropic agents may also affect lysosomal polarity thereby compromising the accuracy of Lyso-OC in sensing autophagy-associated lysosomal polarity changes. Further validation using other means of autophagy induction or inhibition will be necessary to more carefully scrutinize the utilities of Lyso-OC in assessing autophagic flux.
The multistep and dynamic nature of the autophagy process makes it challenging to monitor and quantify. There is a growing awareness across disciplines for the need to determine whether the process has achieved degradative completion via lysosomal fusion and hydrolysis. This need has spurred the development of tools and methods for monitoring autophagic flux assays that have undergone a considerable evolution in the past few years. For example, recognition of the context-dependency of ARs has led to a dramatic expansion in their discovery and a corresponding expansion in their applications in monitoring autophagic degradation. The immunoblot-based LC3B turnover assay, an early standard in the field, is now accompanied by turnover assays for other members of the MAP1LC3 and GABARAP subfamilies. Monomeric fluorescent-tagged versions of these proteins evolved to tandem fluorescently tagged versions, which have been further adapted to include a fluorescent-tagged internal control for measuring reporter construct expression levels. Innovative new strategies incorporating LIR-containing fluorescent sensors also deserve further attention. To help avoid potential artifacts associated with ectopic expression of tagged reporter constructs all together, and in recognition of autophagy-independent roles for the MAP1LC3 and GABARAP family members [29,52], there has also been a move to incorporate methods that do not rely on these proteins. These strategies, like DQ-BSA, Cyto-ID, Keima, HQO, and Lyso-OC, are particularly useful for screening and/or in live cell contexts, with promise for further advances in this area. While many of these newer methods promise a more reliable evaluation of autophagic flux in mammalian cells, it is important to consider measurements at multiple time points and/or under various experimental conditions. Finally, one factor that remains constant in this rapidly expanding field is the recommendation to always employ multiple complementary assays and markers when monitoring autophagic flux.
autophagy-related 16 like 1
autophagy-related 4B cysteine peptidase
BCL2-interacting protein 3
bovine serum albumin
enhanced green fluorescent protein
enzyme-linked immunosorbent assay
- FAM134B, RETREG1
reticulophagy regulator 1
GABA type A receptor-associated protein
GABA type A receptor-associated protein like 1
GABA type A receptor-associated protein like 2
green fluorescent protein
LC3B with C-terminal glycine residue deleted
microtubule-associated proteins 1A/1B light chain 3
microtubule-associated proteins 1A/1B light chain 3 alpha
microtubule-associated proteins 1A/1B light chain 3 beta
microtubule-associated proteins 1A/1B light chain 3 gamma
monomeric red fluorescent protein
Neighbor of BRCA1
- NIX, BNIP3L
BCL2-interacting protein 3 like
N-ethylmaleimide-sensitive factor attachment protein receptor
tandem fluorescent-tagged LC3B (mRFP–GFP–LC3B)
All authors contributed to the writing and editing of this manuscript. All authors have read and approved the final manuscript.
The authors are grateful for support from the Natural Sciences and Engineering Research Council of Canada (NSERC) [grant RGPIN/04982-2015] and the Canadian Institute of Health Research (CIHR) [grant MOP-78882]. K.C.Y. is financially supported by the 2017 Neuroendocrine Tumor Research Foundation — AACR Grant [grant no. 17-60-33-GORS].
The authors thank S. Bortnik, R. Camfield, C. Choutka and J. Xu for helpful discussions and comments on the manuscript.
The Authors declare that there are no competing interests associated with the manuscript.
These authors contributed equally to this work.