Common Assays in Mammalian Golgi Studies

Jie Li; Jianchao Zhang; Sarah Bui; Erpan Ahat; Divya Kolli; Whitney Reid; Lijuan Xing; Yanzhuang Wang

doi:10.1007/978-1-0716-2639-9_20

. Author manuscript; available in PMC: 2023 May 31.

Published in final edited form as: Methods Mol Biol. 2023;2557:303–332. doi: 10.1007/978-1-0716-2639-9_20

Common Assays in Mammalian Golgi Studies

Jie Li ^1,^#, Jianchao Zhang ^1,^#, Sarah Bui ^1,^#, Erpan Ahat ¹, Divya Kolli ¹, Whitney Reid ¹, Lijuan Xing ¹, Yanzhuang Wang ^1,²

PMCID: PMC10231433 NIHMSID: NIHMS1894402 PMID: 36512224

Abstract

The Golgi is a complex structure characterized by stacks of tightly aligned flat cisternae. In mammalian cells, Golgi stacks often concentrate in the perinuclear region and link together to form a ribbon. This structure is dynamic to accommodate continuous cargo flow in and out of the Golgi in both directions and undergoes morphological changes under physiological and pathological conditions. The fine, stacked Golgi structure makes it difficult to study by conventional light or even super-resolution microscopy. Furthermore, efforts to understand how Golgi structural dynamics impact cellular processes have been slow because of the knowledge gap in the protein machinery that maintains the complex and dynamic Golgi structure. In this method article, we list the common assays used in our research to help new and established researchers select the most appropriate method to properly address their questions.

Keywords: Golgi biology, Morphology, Function, Transfection, Viral system, RNA interference, Microscopy, Trafficking, Glycosylation, Fractionation, In vitro techniques, Proteomics, Secretomics, RNA-seq, Proximity labeling

1. Introduction

The Golgi apparatus was firstly described in 1898 by the Italian physician and cytologist Camillo Golgi as a network structure in the cell and was initially named as internal reticular apparatus [1]. The structure of the Golgi was not recognized as a novel cellular organelle until the mid-1950s after the invention of electron microscopy (EM). Designation of the “Golgi complex” entered officially in the literature in 1956 [2]. Ever since, the studies on Golgi structure and function have been tightly related to the development of modern cell biology and biological chemistry.

The Golgi apparatus is now recognized as a well-organized complex consisting of stacks of several parallelly aligned flat cisternae, which are further laterally linked into a ribbon-like structure localized in the perinuclear region. Each stack, the functional unit of the Golgi, can be divided into cis-, medial-, and trans-Golgi sub-compartments, with the cis-side receiving vesicles from the endoplasmic reticulum (ER) and trans-side exporting vesicles to the downstream compartments in the trafficking pathway. Protein processing and trafficking are major functions of the Golgi. While passing through the Golgi stack, cargos go through a series of sequential modifications by a set of enzymes residing in different compartments of the Golgi membranes. The processed proteins are then sorted and passed on to the downstream compartments to their final destination following the secretory pathway.

The development of imaging techniques, including EM and immunofluorescence microscopy, has largely accelerated research on Golgi morphology and structure formation. Several assays can be applied to track the cargo proteins as they travel through the trafficking pathway. Using retention and synchronized release of a fluorescently labeled cargo protein, real-time tracking of the protein trafficking is now possible. Since certain processing steps happen in each compartment of the trafficking pathway where the responsible enzymes reside, protein trafficking can also be indicated by analyzing the processing status of the cargos.

A number of biochemical approaches also facilitated Golgi research. Researchers can investigate the role of specific proteins in Golgi morphology and functions by manipulating the protein level using gene overexpression and interference methods. Additionally, subcellular fractionation methods make it possible to isolate Golgi membranes, and therefore, a cell-free system can be developed to reconstruct the Golgi membrane organization processes. This simplified in vitro system can be a powerful tool when determining the key factors in Golgi structure regulation.

To discover novel factors involved in Golgi structure and function regulation, systems approaches (or omics) for large-scale analysis have recently been developed. When combined with morphology and function assays in manipulated cell lines and controlled experimental conditions, large-scale analysis can identify candidate proteins involved in Golgi morphology and function regulation, as well as new interactions between candidate proteins.

In this article, we summarize the common assays frequently used in Golgi structure- and function-related studies, aiming to give primary guidance to researchers with research interests involving the Golgi apparatus to select the appropriate methods to address their questions.

2. Materials

2.1. Common Reagents

Common reagents used in mammalian Golgi studies are listed in Table 1.

Table 1. Common reagents used in mammalian Golgi studies.

Description	Manufacture	Cat. #
Cell culture
DMEM (Dulbecco’s Modified Eagle Medium)	Invitrogen	11995065
Hyclone bovine calf serum, iron supplemented	GE Healthcare	SH30072.03
Non-viral and viral transfection
Gibco Opti-MEM^™ Medium	Invitrogen	31985070
Polyethylenimine (PEI)	Polysciences	23966–2
Lipofectamine^™ 2000 transfection reagent	Invitrogen	11668019
Lipofectamine^™ RNAiMAX transfection reagent	Invitrogen	13778150
Polybrene transfection reagent	Millipore	TR-1003-G
Cytiva Whatman^™ Uniflo 0.45 μm polypropylene (PES) syringe filter	Cytiva	09-928-063
Immunofluorescence imaging
Paraformaldehyde	Fisher	AC416780030
Albumin, bovine fraction V	Dot	DSA30075–100
1–1/2 Micro-coverglass—12 mm diameter	Fisher	12–545-81
Trafficking assays
Brefeldin A, from Eupenicillium brefeldianum	AG Scientific	B1009
Cycloheximide	AG Scientific	C1189
Sulfo-NHS-SS-Biotin	Fisher	PI21331
Endoglycosidase H (endo H)	New England BioLabs	P0702S
Subcellular fractionation and in vitro reconstitution
Sucrose	American Bioanalytic	AB01900–0100
HeLa S3 cell line	ATCC	ATCC CCL-2
Nocodazole	Fisher	AC358240500
Proteomic and secretomic analysis
Pierce^™ RIPA buffer	Fisher	89900
Amicon ultra-15 centrifugal filter unit, 3KDa	Millipore	UFC900324
TMT10plex^™ Isobaric label reagent set	Fisher	90110

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This table provides examples of reagents and commercial suppliers. Comparable reagents can be obtained from other sources

3. Methods

3.1. Protein Expression, Depletion, and Turnover Assay

To investigate the function of a protein of interest, it is necessary to deplete and/or overexpress this protein. In some cases, transient transfection is sufficient to obtain useful information; in other cases, establishing a cell line that stably expresses the protein of interest provides more convincing and reproducible results. In this section, we introduce non-viral and viral transfection methods that we routinely use to manipulate the expression of proteins. We also include the cycloheximide chase experiment that is broadly used to test protein turnover.

3.1.1. Protein Expression by Transient Transfection

Transfection is a process that introduces (normally using non-viral methods) exogenous DNA into eukaryotic cells for the expression of a gene of interest [3]. Transfection has wide applications in various model organisms and diseases. Transfection, in general, can be classified into transient transfection, where exogenous DNA is not integrated into the genome of the host cell, and stable transfection, in which exogenous DNA is integrated into the host genome. Transient transfection usually does not lead to the integration of the plasmid DNA into the host cell genome, which limits its application to the study of a short-term expression of a gene of interest in the cell [3]. Transient transfection often generates a heterogeneous cell population that expresses the target protein at different levels. While this allows the researcher to detect dose-dependent effects, it can also generate artifacts caused by over-expression. For microscopy, we normally select cells that express the exogenous protein at a modest level; i.e., the exogenous protein is localized in the expected localization in cells by microscopy and is no more than threefold of the endogenous protein by western blot. After transfection, stable expressions can be achieved by long-term drug selection if the transfected plasmid carries a gene that is resistant to a drug (e.g., puromycin) or by picking up single colonies to generate stable cell lines [4], but it is time-consuming and the efficiency is relatively low.

Transient transfection can be achieved using chemical or non-chemical (physical) methods. Chemical-based methods are usually highly efficient, applicable for many cell types, easy to practice, economical, and thus are commonly used in cell biology research. To express proteins of interest using chemical-based methods, a transfection complex is prepared by mixing plasmids encoding the gene(s) of interest and transfection reagents, such as polyethylenimine (PEI) or lipofectamine in serum-free medium (e.g., Opti-MEM) according to the manufacturer’s instruction, and then add it into the culture medium. In chemical-based transfection, transfected cells are usually collected at 24–48 h post-transfection to analyze the expression efficiency and downstream effects [5]. Non-chemical-based transfection uses physical forces such as electroporation or injection of DNA into the cells.

3.1.2. Establishment of Stable Cell Lines Using Viral Systems

In contrast to transient transfection, the use of viral vectors can stably express the gene of interest by integrating the exogenous DNA into the host genome (Table 2). Two viral transduction systems commonly used to establish stable expression cell lines are retrovirus and lentivirus [6–8]. Lentiviruses are ideal for experiments that require permanent changes in desired cell lines due to their ability to infect both dividing and non-dividing cells [9]. On the other hand, retroviruses can only infect dividing cells. Other viral vectors such as adenovirus have high transduction efficiency, especially toward primary cells; however, the expression is only transient. Viral vectors are typically chosen to suit the packaging needs, duration of expression, and transduction efficiency.

Table 2. Comparison of non-viral and viral systems in protein expression.

	Non-viral		Viral
	Liposome	Molecular conjugate	γ-retrovirus	Lentivirus
Genome	DNA plus lipid	DNA plus polypeptide	ssRNA (7–20 kb)	ssRNA (9 kb)
Packaging capacity	>10 kb	>10 kb	6–10 kb	8–9 kb
Transduction	Dividing/non-dividing cells	Dividing/non-dividing cells	Dividing cells	Dividing/non-dividing cells
Transduction efficiency	Low	Low	Moderate	Moderate
Expression	Transient	Transient	Stable	Stable
Integration	No	No	Integrating	Integrating
Biosafety level	BSL-1	BSL-1	BSL-2	BSL-2
Immunogenicity	Low	Low	Moderate–high	Moderate–high
Gene therapy strategy	Ex vivo	Ex vivo	Ex vivo	Ex vivo

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For safety precautions, viral transduction systems separate the necessary components to package an infectious virion into multiple different vectors: packaging plasmid, envelope plasmid, and transfer plasmid. The packaging plasmid usually encodes the viral enzymes (gag-pol). The interchangeable envelope plasmid determines the infectivity range of the virus, which is referred to as the tropism of the virus pseudotype. Vesicular stomatitis virus G (VSV-G) envelope protein is most commonly used to render the virus amphotropic (capable of infecting mammalian cells). Lastly, the transfer vector contains the gene of interest. These three plasmids are co-transfected (using a transfection reagent like FuGENE) into a packaging cell line, typically HEK293T (see Note 1), which will produce infectious viral particles released to the supernatant that can be harvested to infect target cells. Collection of viral supernatant can begin 48 h post-transfection up to 96 h post-transfection, although titers may decrease over time. The viral supernatant is filtered through a 0.45 μm filter (see Note 2) and can be concentrated via ultracentrifugation if desired (see Note 3). Optimization is necessary to determine the best transduction conditions (see Note 4). The initial transduction yields a polyclonal cell population, where the integration varies from cell to cell because the integration event is random. There is also a different number of integration events from cell to cell, particularly if the multiplicity of infection (MOI) is greater than one. To select for cells with similar integration sites and events, monoclonal colonies can be selected for expansion (see Note 5). Stable cell lines established via viral transduction offer greater reproducibility in long-term studies.

3.1.3. Gene Knockdown

There are a few methods to reduce gene expression (knockdown, KD), including small interfering RNA (siRNA), short hairpin RNA (shRNA), antisense oligonucleotides (ASO), and CRISPR-based gene knockdown (CRISPRi) [10, 11]. Among these, siRNA-based KD is frequently used in mammalian cells [12]. In gene KD using siRNA, siRNA complementary to the target gene is exogenously introduced into the cell to trigger degradation of the target mRNA or inhibits its translation. A transfection complex is prepared by mixing siRNAs and transfection reagent RNAiMAX (Invitrogen) in Opti-MEM according to the manufacturer’s instruction and then added into the culture medium. Cells are usually collected after 48–96 h post-transfection to analyze the KD efficiency (see Note 6). It is generally believed that gene KD may cause side effects. Therefore, multiple siRNA oligos are routinely used (separately or pooled) for the same gene, while rescue by expression of a siRNA-resistant construct is now a standard method to rule out off-target effects.

3.1.4. Cycloheximide (CHX) Treatment to Determine Protein Turnover

The protein abundance in a cell is controlled by synthesis (transcription and translation) and degradation (proteasomal and lysosomal). The most commonly used technique in analyzing protein turnover kinetics in early years required radioactive isotope pulse-chase labeling, immunoprecipitation, and autoradiography. A convenient alternative approach to studying protein stability is to perform a CHX chase experiment. This technique is suitable for studying the degradative fate of a wide range of proteins that have relatively short half-lives [13].

CHX blocks protein synthesis by inhibiting translation elongation [14–16]. It is a cell-permeable molecule and therefore can be directly added to cell culture medium. CHX is typically used at 10–100 μg/mL for 4–24 h (see Note 7). Cells are collected and immediately lysed at the desired time points of the treatment. Lysates are typically separated by polyacrylamide gels electrophoresis followed by western blot [17, 18]. Moreover, CHX is frequently added in protein trafficking assays to track the pre-existing cargo proteins and avoid the noise caused by newly synthesized proteins [19]. A combination of CHX with proteasomal and lysosomal inhibitors can provide useful information on how the target protein is degraded. As demonstrated by the wide range of assays compatible with cycloheximide treatment, this technique is therefore a highly versatile method in Golgi biology.

3.2. Imaging Techniques for Morphological Studies

The Golgi is part of the endomembrane system, characterized by its unique structure formed by a series of stacked flat cisternae that are laterally linked to form the Golgi ribbon. Being a well-organized, yet dynamic organelle, Golgi morphology is tightly linked to its function. Therefore, imaging techniques represent an essential approach in studying Golgi structure formation, dynamics, and function.

3.2.1. Immunofluorescence Microscopy

Immunofluorescence (IF) microscopy is an important immunochemical technique widely used in cell biology research. It can be used to determine a broad range of Golgi properties, including the detection of Golgi protein localization [19], Golgi morphology [9], protein trafficking [20], and glycosylation [21]. By combining the usage of specific antibodies and fluorophores, IF facilitates the visualization of a wide variety of antigens in various cell types and preparations. Generally, two IF methods can be employed: direct (primary) IF or indirect (secondary) IF. Both methods use antibodies to recognize specific antigens. The difference is that the former uses fluorophore-conjugated primary antibodies, and so only one antibody is sufficient to detect one antigen, while the latter uses unlabeled primary antibodies and fluorescently labeled secondary antibodies. Due to its high sensitivity, signal amplification, and ability to detect several components spontaneously, indirect IF is more widely used.

A standard IF protocol involves two steps of incubation: first, a primary antibody binds to the target epitope in cells; second, a fluorophore-tagged secondary antibody recognizes and binds to the primary antibody [22]. In most cases, cell membranes need to be permeabilized by detergent to allow the antibodies to enter the cell; in some other cases, cell surface staining without permeabilization allows the detection of proteins only at the exterior surface of the cell. In the past decades, the development of advanced fluorescence microscopy techniques, including but not limited to fluorescence recovery after photobleaching (FRAP), Förster or fluorescence resonance energy transfer (FRET), fluorescence life-time imaging microscopy (FLIM), and super-resolution microscopy, has significantly broadened the usage of IF, improved image qualities, and made quantitative microscopy possible [23, 24]. In addition, the cloning and invention of a series of fluorescent proteins and probes further enhanced the capability of fluorescence microscopy.

3.2.2. Electron Microscopy (EM)

EM is a widely used technique to obtain high-resolution images of biological and non-biological specimens. There are two main types of electron microscopy: transmission EM (TEM) and scanning EM (SEM). SEM produces images of a sample by scanning the surface with a focused beam of electrons. For TEM, a beam of electrons passes through a thin sample (usually below 100 nm) to produce an image, so the sample’s thickness and composition play a significant role. TEM is more widely used by biologists to investigate fine structures in tissues, cells, subcellular membrane organelles, and organelle contact sites.

In 1954, the first TEM images of the Golgi apparatus were shown as a stack of curved smooth-surface cisternae [25, 26], which was a milestone in Golgi research because it resolved the controversy regarding whether Golgi’s apparatus is a real structure or an artifact [27]. Although fluorescence microscopy is more broadly used in current research, EM provides high-resolution images with more details that IF still cannot match. Immuno-EM, an extended TEM method, can label different Golgi markers at ultrastructural levels and has revealed small soluble proteins’ movement through the Golgi by diffusion (see Note 8) [28]. Currently, EM is an essential tool to study the ultrastructure of the Golgi apparatus in vitro, in cells, as well as in tissues [29, 30].

3.2.3. EM Tomography

EM tomography (also referred to as electron tomography, ET), extended from traditional TEM, is a common technique for obtaining detailed 3D structures. ET reconstructs the complete 3D structure from a series of two-dimensional images [31]. Alignment and digital combination of all projections result in an accurate representation of the original object. ET obtains 3D structural information of cells and organelles at resolutions that range between 2 and 10 nm. The Golgi apparatus in mammalian cells is a complex and dynamic organelle displaying interconnected highly organized stacks of cisternae. ET can provide good insights into the complex Golgi architecture and fine structure–function relationships. 3D Golgi structures by ET in normal rat kidney (NRK) cells revealed that most vesicular profiles visualized in the trans-Golgi network (TGN) are connected to TGN tubules and that TGN tubules extended from the margins of both cis- and transcisternae, suggesting that these tubules contribute to traffic to and/or from the Golgi, providing extensive novel insights into the structure and function relationship [32, 33]. Recently, whole-cell in situ cryo-ET of various SARS-CoV-2-infected cell lines revealed the double-membrane vesicles (DMV) morphology, virus assembly site, and extracellular virions close to their native state [34]. Also, a combination of electron tomography and focused ion beam scanning electron microscopy (FIB-SEM) showed fragmented Golgi cisternae in the vicinity of double-membrane vesicles (DMVs) in SARS-CoV-2 infected Calu-3 cells [35].

3.3. Common Assays to Study Golgi-Mediated Trafficking

As the transport center in the cell, a major function of the Golgi is protein trafficking. Tracking cargo proteins as they travel through the Golgi can be challenging. Here, we introduce a few easy-to-use assays that can be used to monitor protein trafficking through the Golgi.

3.3.1. Brefeldin A Treatments and Washout

Brefeldin A (BFA) treatment is known to reversibly block ER-to-Golgi transport by inhibiting GBF1, a guanine nucleotide exchange factor (GEF) of ARF1, which prevents the formation of COPI vesicles [36]. BFA blockage and release are widely used in protein trafficking studies. Generally, treatment of cultured cells with 0.25–5 μg/mL BFA for 0.5–2 h, depending on the cell line and marker used, is sufficient to cause the absorption of Golgi membranes into the ER. Washout of BFA leads to the release of proteins from the ER that then synchronously traffic to the Golgi. Combining with different experimental manipulations such as knockdown of a trafficking-related protein, expression of a mutant, or inhibition of protein synthesis by CHX treatment allows for determining their roles in ER-to-Golgi trafficking (see Note 9). By fluorescence microscopy or subcellular fractionation, the trafficking of a cargo molecule can be visualized or detected at different time points after the release. When samples are analyzed by western blot, one can also determine whether a protein of interest is cleaved (e.g., sterol regulatory element-binding protein (SREBP) [37]) and the timeline of the cleavage after the release from the ER and thus estimate where in the cell (e.g., Golgi, TGN, post-Golgi vesicles or lysosomes) the protein is cleaved. BFA is also often used to determine whether a protein is secreted through the conventional secretory pathway.

3.3.2. Temperature Block and Release of VSV-G tsO45

The envelope “G” protein of vesicular stomatitis virus (VSV-G) is a viral glycoprotein transported through the secretory pathway. VSV-G tsO45 is a thermosensitive (ts) mutant of VSV-G. VSV-G tsO45 can be expressed by transient transfection or by adenoviral transduction [38]. When synthesized at the non-permissive temperature of 40.5 °C, VSV-G tsO45 is incompletely glycosylated and misfolded, forms non-covalently associated aggregates, and is retained in the ER [39]. When cells are shifted to the permissive temperature of 32 °C, the aggregates of VSV-G tsO45 disassemble, which can then correctly fold, trimerize, exit ER, and traffic through the Golgi to the plasma membrane [38, 40]. When combined with different experimental conditions, this allows the characterization of a protein of interest in membrane trafficking. By fusing VSV-G tsO45 with fluorescent proteins, its transport from the ER to plasma membrane can be observed in real-time by live cell imaging. Given that VSV-G is N-glycosylated, a combination with endoglycosidase H (endo H) treatment can help with understanding the speed of membrane trafficking (see Subheading 3.3.4). Therefore, it has been one of the most used tools in membrane trafficking studies.

The technique of temperature block and release is frequently but not exclusively used with VSV-G O45. Temperature block and release is also used as a tool to study protein trafficking at distinct stages of the secretory pathway. By adjusting the temperature, secretory proteins can be accumulated in particular compartments in the trafficking pathway and their exit from the compartments can be synchronized. Using membrane proteins as markers, it was reported that incubation of the cells at 15 °C leads to retention of cargo proteins in the early Golgi or ER-Golgi intermediate compartment (ERGIC), while incubation at 20 °C leads to a block at the TGN [41, 42]. For example, one can first block cells at 15 °C and then shift cells to 20 °C to specifically test the speed of trafficking between the cis- and trans- Golgi under different experimental conditions [21]. The exact mechanism behind the temperature block is not fully understood, but may be related to the membrane fluidity [43] and the association of Golgi proteins such as ARF1 and ARL1 affected by the temperature change [44]. Therefore, one major drawback of this technique is that the process exposes the cells to non-physiological temperatures that affect trafficking. Moreover, a rapid and precise switch of temperature (see Note 10) is not always guaranteed and therefore variations among experiment repeats may be introduced.

3.3.3. Synchronized Release of Cargo Proteins Using the Retention Using Selective Hooks (RUSH) System

To overcome the drawback of the temperature block and release, Boncompain and Perez developed the RUSH system, which makes it possible to simultaneously release a pool of secretory proteins from particular compartments (e.g., the ER) and monitor their anterograde trafficking under physiological conditions [45]. This secretory assay is a two-stage method based on the retention of a chosen reporter (a cargo protein) in a donor compartment and the induction of its synchronous release. The retention of the reporter is facilitated by its interaction with a hook protein that stably resides in the donor compartment, such as the ER or Golgi. The interaction is mediated by streptavidin fused to the hook protein and the streptavidin-binding peptide (SBP) fused to the reporter. This interaction is disrupted upon the addition of biotin into the culture medium. Because of its high affinity to streptavidin, biotin competes out the SBP-fused reporter and thus enables the release of the reporter to its downstream compartment in the trafficking pathway (see Note 9) [46].

Real-time visualization of the reporter is possible when a fluorescent protein is fused to it. Combining the non-permeabilization process and antibodies recognizing their extracellular domains, the plasma membrane proteins can be detected using IF or flow cytometry analysis to indicate the trafficking rate [47]. The RUSH system is an ideal alternative for the traditional temperature block and release technique (see Note 11), since simply adding biotin into the medium is easier to perform than rapid and precise temperature switching; it also avoids possible artifacts caused by the non-physiological temperature treatment.

3.3.4. Pulse-Chase Labeling and Endo H Treatment

In the early years, radioactive isotope pulse-chase labeling was used to monitor protein trafficking by many researchers. To increase the resolution of trafficking, pulse-chase labeling is often combined with endo H treatment. Endo H is a glycosidase that removes high-mannose oligosaccharides but not complex oligosaccharides from target proteins [48]. N-linked protein glycosylation initiates in the ER with the synthesis and transfer of a high-mannose oligosaccharide precursor to nascent proteins. Upon transport to the Golgi, N-glycans are further processed by Golgi-resident glycosidases and glycosyltransferases that remove mannoses and add other sugar residues such as galactose, GlcNAc, and sialic acid [49]. Because of the different sugar compositions in N-glycans attached to membrane and secretory proteins, the ER and cis- Golgi forms of these proteins are sensitive to endo H treatment, while they become resistant to endo H treatment once arrive the medial-/late Golgi. When analyzed by SDS-PAGE and autoradiography, a shift of the band indicates that the protein is in the ER or cis-Golgi, and a lack of band-shift suggests that it has reached late Golgi [21]. In many times, peptide-N-glycosidase F (PNGase F) is used as a control as it removes all types of N-glycans from core proteins regardless of their sugar compositions; this ensures that the band-shift assay works.

While radioactive isotope pulse-chase labeling is less commonly used in current research, endo H is still widely used, most often in combination with “trafficking retention and release” assays such as VSV-G tsO45 and the RUSH assay introduced above. Briefly, a target secretory protein is trapped in the ER and released for secretion in the presence of CHX for certain periods (chase) (see Note 12). The cells are collected at different time points, and cell lysates are treated with endo H (or PNGase F). The molecular weight shift of the protein shown on western blot due to deglycosylation is used to reveal when the protein has reached the medial- Golgi [9, 50]. In addition, endo H can be used to test whether a membrane or lumenal Golgi protein resides in cis- or medial/trans- Golgi if this protein is glycosylated. As N-glycosylation occurs only in the lumen of the secretory pathway, a protein that is sensitive to endo H or PNGase F is normally believed to be a membrane or secretory protein.

3.4. Subcellular Fractionation and In Vitro Reconstitution Using Isolated Golgi Membranes

Subcellular fractionation is an important approach in Golgi studies, which allows the determination of biochemical compositions of cellular samples, including proteins, lipids, and oligosaccharides. The results can be used to interpret the structural organization and function of the Golgi. To investigate the key factors affecting Golgi membrane organization, in vitro assays using isolated Golgi membranes were developed. By incubating Golgi membranes with prepared cytosol or purified proteins, alterations in Golgi organization can be observed in a simplified cell-free system. Several factors involved in Golgi biogenesis and morphology regulation have been identified using such assays [29].

3.4.1. Differential Centrifugation to Separate Different Cellular Structure

Differential centrifugation is a widely used method for the separation of various organelles based on the sedimentation rates. Differential centrifugation achieves separation of organelles by a stepwise increasing speed. Low speed (usually <1000 g) centrifugation sediments unbroken cells and nucleus, and the speed is then increased until the target membranes are pelleted. Generally, the plasma membrane, nuclei, and mitochondria pellet first, mitochondria may also appear in the next pellet along with lysosomes and peroxisomes, then Golgi apparatus and microsomes in the following pellets.

Differential centrifugation was first introduced in 1934 in order to obtain high purity of mitochondria [51] and later adopted to separate Golgi membranes from vesicles. Tissues (usually rat livers) were homogenized in a homogenization buffer (usually 0.5 M sucrose, 5 mM MgCl₂, 0.1 M phosphate buffer pH 6.7, protease inhibitor cocktail) by gently pressing through a 150 μm mesh stainless steel sieve [52–56]. The Golgi membranes pellet under a low centrifugation speed of 15,000–30,000 g for 10–20 min depending on the homogenization method and buffer used, while the vesicles remain in the supernatant [57]. Vesicles will be pelleted after ultracentrifugation of 100,000 g for 60 min [20]. Protein levels from each separated fraction could be easily determined by western blot [47]. Activity assays of enzymes concentrated in the Golgi or other subcellular compartments can be performed to confirm the purity. The purity and morphology of target membranes can also be checked by EM [29]. As these fractions are not pure, gradient centrifugation may be applied to further fractionate or purify target membranes [58].

3.4.2. Equilibrium Gradients to Fractionate Membrane Organelles

Equilibrium density gradient centrifugation is a method used to separate cellular membrane structures based on density. During gradient centrifugation, the subcellular structures in the sample will travel to a zone in the medium’s density gradient that is equal to the structure’s density. The density of a membrane organelle depends on its composition. As proteins are heavier than lipids, the more proteins a membrane structure contains relative to lipids, the denser this membrane structure is, and the deeper it goes in the equilibrium density gradient. For example, secretory granules contain a high protein content and thus are often found in deeper fractions of the gradient. In general, mitochondria are heavier than the Golgi, while ER microsomes could be found in a variety of fractions depending on the number of ribosomes associated with them. A commonly used medium in the gradient is sucrose, and both continuous and step gradients can be used to separate cellular membrane structures [56, 59]. In our Golgi purification procedure described below (see Subheading 3.4.3), the Golgi is highly enriched at the 0.5/0.86 M sucrose interface [29, 55, 56].

In 1951, an equilibrium gradient was introduced to separate a potato yellow dwarf virus [60]. Later, this method was applied to isolate the Golgi membranes from mammalian liver [61]. Additionally, the separation of Golgi-derived vesicles from Golgi remnants in the Golgi disassembly and reassembly assays can be accomplished using the equilibrium gradient centrifugation [29]. Western blot, EM, or enzymatic activity analyses of enzymes residing in the Golgi and other organelles of interest can be performed to confirm the purity [62].

3.4.3. Purification of Golgi Membranes from Animal Tissues

Purification of Golgi membranes from animal tissues yields an abundant source of material, which can be used for structural observation under physiological conditions and in vitro functional assays. The zonal centrifugation methods for Golgi membrane preparation from bovine [61] and rat livers [52, 53] were first developed in the 1970s. Mammalian livers have abundant Golgi membranes to satisfy the high demand of protein secretion, which makes them a perfect source for Golgi membrane preparation. The Golgi membrane purification by two sequential sucrose gradients was developed, and the purity was significantly increased [55, 63]. The first gradient (0.25–0.86 M sucrose) separates intact cells and the majority of cytosol out of Golgi membranes (see Note 13). The second gradient (0.5–1.3 M sucrose) further concentrates the Golgi membranes by separating cytosol and other membrane organelles [29, 55, 56].

The yield of the Golgi membranes can be determined by calculating the ratio of the activity of a Golgi enzyme, β−1,4-galactosyltransferase (GalT) in purified Golgi membrane over that of the total liver homogenate. Purified Golgi membranes are usually 80- to 100-fold enriched over the homogenate and 60–70% of Golgi membranes form stacks, which can be confirmed by EM [29]. Purified Golgi membranes can be used for analyzing the Golgi localization of target proteins by western blot or in vitro reconstitution of the Golgi disassembly and reassembly process during cell division, as described below (see Subheading 3.4.4).

3.4.4. In Vitro Budding Assay

Protein trafficking through the Golgi could be explained by two major models, cisternal maturation and vesicular transport [64]. The maturation model proposes that cargos are transported within the cisternae by the movement of the cisternae, and Golgi-resident enzymes are recycled via retrograde transport of COPI vesicles. In the vesicular transport model, Golgi-resident proteins, including glycosylation enzymes, are retained in the cisternae, while cargos are successively transported through all cisternae by COPI vesicles. In both models, the budding rate of vesicles determines the rate of transport across the Golgi [65]. We have developed an in vitro budding assay by incubation of purified Golgi membranes with AFR1 and coatomer to determine the content of COPI vesicles. Essentially, when purified Golgi membranes are incubated with purified AFR1 and coatomer (with or without purified mitotic kinases to mimic mitotic conditions), or interphase cytosol as an alternative, Golgi membranes are fragmented into COPI vesicles [66, 67]. Golgi remnants and COPI vesicles can then be separated by equilibrium centrifugations as described above and analyzed by western blot [29, 68, 69] or proteomic approaches [70]. Our results showed that Golgi enzymes are enriched in COPI vesicles [66, 67] and thus support the cisternal maturation model.

The in vitro budding assay has also been a useful tool to identify new factors that are involved in Golgi membrane fusion and protein trafficking, such as the N-ethylmaleimide-sensitive factor (NSF) [71, 72] and soluble NSF attachment proteins (SNAPs) [73]. It has also been employed to determine whether unstacked single cisternae can form COPI vesicles more efficiently than stacks [20]. When Golgi membranes were treated with the mitotic kinases CDK1 and Plk1 to phosphorylate GRASP65 and thus unstack the Golgi cisternae, unstacked Golgi membranes showed a higher COPI budding rate than stacked Golgi membranes [68, 69], supporting our conclusion that Golgi stack formation slows down protein trafficking to ensure accurate protein glycosylation and sorting [9, 20, 21].

3.4.5. In Vitro Reconstitution of Golgi Disassembly and Reassembly During Cell Division

Similar to the in vitro budding assay described above (see Subheading 3.4.4), a stepwise treatments of purified rat liver Golgi membranes with mitotic and interphase cytosol allow the disassembly and reassembly of the Golgi structure as seen during cell division [74]. Interphase and mitotic cytosols are prepared from HeLa S3 cells. For mitotic cytosol, cells are blocked to G2/M phase by nocodazole treatment (see Note 14). When Golgi stacks were incubated with mitotic cytosol (MC), they were transformed into vesicles and short tubules. When these mitotic Golgi fragments (MGFs) were reisolated by centrifugation and further incubated with interphase cytosol (IC), they fused to form new cisternae that subsequently formed stacks. Following this, a modified form of this in vitro membrane fusion assay was developed, in which instead of cytosol, purified recombinant proteins are used to treat Golgi membranes [29]. The modified assay using purified proteins helped reveal the minimal machinery required for Golgi disassembly and reassembly during the cell cycle. In addition, the cell-free in vitro fusion assay also allowed the identification and characterization of a number of proteins involved in the cell cycle-regulated Golgi disassembly and reassembly process, including GRASP65 [75, 76], GRASP55 [77], and p115 [78].

3.4.6. In Vitro Reconstitution of Golgi Structure Formation and Function in Semi-Permeabilized Cells

The cholesterol content of the plasma membrane is much greater than that of intracellular membranes. The semi-permeabilization assay utilizes this property to treat cells with digitonin, a detergent that reacts specifically with cholesterol and thus selectively disrupts the plasma membrane but not intracellular membrane structures [79]. Taking advantage of semi-permeabilization which maintains the integrity of cellular organelles such as the ER and Golgi, Balch and coworkers used this system to reconstitute ER-to-Golgi trafficking in vitro by incubating semi-permeabilized cells with cytosol and ATP [80]. This assay was based on the sensitivity of the Man_5-GlcNAc₂ oligosaccharide form of the VSV-G protein to the enzyme endoglycosidase D (endo D). VSV-G tsO45 was used due to its sensitivity to the restrictive temperature. Under low temperature, the VSV-G tsO45 protein was transported from the ER to the Golgi marked by the presence of the Man₅GlcNAc₂, an endo D-sensitive form [79].

In addition to digitonin, semi-permeabilization can be induced by freeze–thaw, ilimaquinone, or a bacterial toxin streptolysin O [81]. Using the in vitro semi-permeabilization assay, it was successfully shown that post-mitotic Golgi structure formation depends on NSF and an NSF-like AAA ATPase, p97 [82]. Since the plasma membrane is selectively disrupted and permeabilized to chemical molecules and antibodies, this semi-permeabilization assay is an excellent model for reconstitution of various cellular processes.

3.5. Systems Approaches in Golgi Studies

With the aim to discover novel factors involved in Golgi structure and function regulation, large-scale analysis is used. Combining these assays with manipulated cell lines and/or experimental conditions, the functions of certain proteins in membrane trafficking and potential binding partners of a protein of interest can be identified.

3.5.1. Proteomic and Secretomic Analysis

Proteomic and secretomic approaches are often used to characterize the entirety of proteins in cells and conditioned media, respectively. The term “proteomics” was first introduced in 1995 as a large-scale analysis of proteins [83]. The major steps in proteomics include sample preparation, protein digestion and tandem mass tag (TMT) labeling, liquid chromatography–mass spectrometry analysis (LC-multinotch MS3), and data analysis. Take the study of the global effect of GRASP55 depletion on protein synthesis, stability and secretion as an example, the procedure for the proteomic and secretomic analyses is depicted in Fig. 1 [5, 84, 85]. In order to avoid the interference of bovine serum albumin (BSA), the secretome sample is generally prepared in serum-free medium for 4–24 h. If an exosome fraction is desired, the medium can be ultracentrifugated at 120,000 g for 90 min at 4 °C. The final concentration of the lysate and medium samples are critical for the quality of MS analysis since low concentration (<1 mg/mL) leads to loss of low expressing proteins in labeling and analysis.

Fig. 1 — Procedure of sample preparation for proteomic and secretomic analysis. Three replicates of each WT and 55KO cells were cultured to 80% confluency. Cells and conditioned medium were collected. The cells were washed with PBS and lysed in Pierce^™ RIPA buffer (Thermo, Cat# 89900). The collected conditioned media were cleaned by two sets of centrifugations, filtered and concentrated with a 3 kDa cutoff filter (Millipore, Cat# UFC900324). The protein concentration of lysate and media samples was tested with Bradford assay and normalized. The samples were trypsinized, TMT-labeled following the manufacturer’s protocol (Thermo Fisher, Cat # 90110), processed in LC-multinotch MS3, and the data were analyzed with Proteome Discoverer (v2.4; Thermo Fisher)

3.5.2. RNA-seq and Transcriptomic Analysis

Transcriptomics is the study of the full set of transcripts within a cell and is a common technique, often used in conjunction with quantification, to profile the differential expression of cells in various phenotypes, environments, or stages of development. Hybridization microarrays and RNA-seq are two common methods for generating the transcriptome. RNA-seq is performed through next-generation or Illumina sequencing and is followed by mapping to a reference library for the organism of interest, alignment, and quantification. A more thorough pipeline for performing RNA-seq is shown in Fig. 2. RNA-seq can be used to better understand the effects of Golgi stress on the cell and may highlight potential relationships between differential protein processing in the Golgi and effects on gene transcription [84].

Fig. 2 — RNA-seq procedure. RNA-seq is performed through next-generation or Illumina sequencing followed by mapping to a reference library for the organism of interest, alignment, and quantification

Gene ontology (GO) is a vocabulary used to describe groups of genes and gene products that serve in a conserved biological process, molecular function, or cellular component among various organisms. This system was developed by The Gene Ontology Consortium and first published in 2000 utilizing the genome information from mouse, Saccharomyces, and Drosophila databases [86]. Since then, the gene ontology system has grown considerably with the increase in available genomic information from a much wider range of organisms. Now, gene ontology is a commonly paired method with omics-based approaches to cluster large datasets of genes or gene products into distinct ontologies to summarize and elucidate differences in biological or molecular functions between cells. Methods used to analyze RNA-seq data, including GO and other similar enrichment or clustering methods, are described in Table 3 as well as commonly used visualization tools summarized in Table 4.

Table 3. Tools and databases for analyzing RNA-seq data.

Tool	Description
GSEA [98] (see Note 15)	Calculates a normalized enrichment score quantifying the overrepresentation of sets of genes in normalized ranked gene sets representing different phenotypes. GenePattern modules also include Bowtie, Tophat, RNA-SeQC, among others.
Enrichr [99, 100] (see Note 16)	Produces a ranked list of enrichment terms determined by degree of overlap between quantitative or crisp gene sets and the unranked input data.
Revigo [101] (see Note 17)	Summarizes gene ontologies through semantic similarity and visualizes clusters of similar ontologies to emphasize the most significant sets of ontologies in a large dataset.
AmiGO [86, 102]	Database to search the gene ontology and annotated genes from other associated databases. Includes the full gene set and GO hierarchies for each term. Includes an enrichment tool.
Gorilla [103]	An enrichment tool to determine novel GO terms that are well represented in the top of a p-value ranked gene list or GO terms that are enriched as compared to a background dataset.
CEMiTool [104]	Identifies co-expression modules within a gene dataset and performs enrichment analysis of the modules. Module expression activity can be compared between different samples and co-expression data can be integrated with protein–protein interaction data to develop an interaction network.
KEGG [105]	Database with KEGG pathway maps determined from published literature and an enrichment analysis through KEGG mapping, a system to compare an input list of genes or proteins with KEGG pathway maps and KEGG modules.
DAVID [106]	A functional tool that can identify gene groups based on annotation similarity and ranked by importance. Significant biological processes are then determined based on the similarity of genes between different gene groups.

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Table 4. Visualization tools for analyzing RNA-seq data.

Tool	Description
cirGO [107]	Visualizes annotated gene expression data that are summarized to remove redundancy through a similarity analysis and grouped by hierarchical clustering. The two-level circular plot includes grouping of annotation terms in the inner ring and non-redundant annotations in the outer ring ranked by highest enrichment. Utilizes Revigo for initial GO term summarization.
String [108–113]	An enrichment tool for visualizing protein–protein interaction networks. Proteins are preferentially mapped into the network based on interaction confidence, creating a network based on the degree of interaction. Network maps can also be annotated based on the evidence for interaction. Co-expression and co-occurrence can also be visualized using this tool.
Tree Map [101]	Visualizes hierarchical structures within the set of GO terms by creating representative rectangular clusters of groups of GO terms which are also grouped into superclusters of related groups. Sizing of the clusters is based on p-value or frequency of GO term as compared to a background dataset. Embedded into Revigo for visualization.
ggplot2 [114]	Visualization tool in programming language R which is highly customizable for aesthetics, formatting, and plot types.
GOBubble [115]	Plots enriched GO terms by p-value and z-score with area proportionate bubbles corresponding to the gene count for each GO term. Bubbles are also colored by clusters of GO terms. GOBubble includes a simple function for removing redundant GO terms. GOBubble is available within the GOplot R package.

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3.5.3. Proximity Labeling Using BioID, TurboID, and Apex

When it comes to large-scale analysis of protein–protein interactions in animal cells, one can think of two broad classes of techniques: (1) fractionation and affinity purification (e.g., immunoprecipitation and pull-down) in which an organelle or protein complex is enriched before its contents are identified by mass spectrometry; (2) imaging and biochemical techniques that rely on antibodies and therefore limit the number of proteins one can detect in a single experiment. One shortcoming of the traditional proteomic workflow that begins with purification of an organelle or protein complex is that cell lysis disrupts compartments and complexes, leading to a loss of key components and increasing chances of contamination. Purification can take a long time during which the proteome can change. But the most important limitation of the traditional mass spectrometry workflow is that many compartments cannot be purified. One example is the sub-compartments of the Golgi stack or the space between Golgi cisternal membranes within the stacks. Some other examples include the mitochondrial intermembrane space, neuronal synaptic cleft, and stress granules. To satisfy this need, proximity labeling assays were developed to bypass the need for purification but still deliver subcellular and spatial information.

The general idea of proximity labeling is to tag proteomes of interest in live cells and subsequently isolate the tagged proteins and identify them by mass spectrometry. The key to this approach is a promiscuous labeling enzyme. One example is using a biotin ligase that utilizes free biotin and converts it into a reactive molecule that then diffuses out of the active site of the enzyme and covalently tags nearby proteins [87]. But this reactive species is unstable and short-lived and therefore will only have a labeling radius of ~10 nm [88, 89]. More distal proteins are not tagged, maintaining the specificity of the interactome of interest. The proteins that are biotinylated can be isolated via streptavidin beads and then identified by mass spectrometry [90].

There are two broad families of enzymes for proximity labeling: peroxidase family and biotin ligase family (Fig. 3). The peroxidases oxidize phenols into radicals for tagging and the biotin ligases convert biotin into adenylate esters for tagging. APEX2 is derived from plant ascorbate and uses hydrogen peroxide to generate a biotin-phenoxyl radical which labels protein tyrosine side chains [91]. Additionally, APEX2 can survive fixation and therefore can be used in conjunction with a DAB (3, 3⁰-diaminobenzidine) reaction to visualize the APEX2 construct subcellular localization at ultraresolution by EM. APEX2 will catalyze the oxidative polymerization of DAB into a locally deposited polymer that recruits electron-dense osmium. The labeling time of APEX2 in cells occurs within 10 s–1 min; the fast-labeling time makes it suitable for experiments in cell culture.

TurboID was engineered by directed evolution from bacteria biotin ligase (BirA). It catalyzes the same chemistry as BioID but is about 100-fold faster. TurboID uses biotin and ATP to generate biotin-AMP which then diffuses out of the active site and covalently tags proteins on their lysine side chains [87]. Due to the non-toxic labeling conditions (no H₂O₂), TurboID is compatible for in vivo as well as in vitro experiments. Greater temporal control is achieved with TurboID, as it has a faster labeling time compared to other biotin ligases such as BirA*, BioID, and BioID2 which can take up to 18 h to biotinylate proteins [92]. Although easy to use, this technique still needs optimization (e.g., biotin concentration, and labeling time) to improve the specificity and labeling efficiency. Applications of proximity labeling to study protein movement, map protein subclasses by location and function, and engineer new strategies for conditional labeling (light, cell-cell contact regulated, calcium ion regulated) are certainly promising and enticing future directions for this molecular tool [93–96].

Acknowledgments

This work was supported by National Institutes of Health (Grant R35GM130331) and the Fast Forward Protein Folding Disease Initiative of the University of Michigan to Y. Wang.

Footnotes

Other packaging cell lines may already contain episomes with the viral machinery and therefore only the transfer plasmid needs to be transfected [e.g., Phoenix cells (Nolan Lab)] [97].

Polyethersulfone (PES) (low-binding) filters are recommended.

For non-VSV-G pseudotyped viruses, ultracentrifugation and repeated freeze–thaw cycles are not recommended, as they are less stable.

⁴

Polybrene can be added to the viral supernatant to facilitate viral particle interaction with target cell membrane. However, polybrene can be toxic to cells; 3–10 μg/mL is a good working range.

⁵

By antibiotic selection or fluorescence-activated cell sorting (FACs) if construct expresses a fluorescent tag.

⁶

Transfection time and KD efficiency may vary depending on the siRNA targeting sequence, concentration of siRNAs, cell type, transfection reagent, and time. siRNA, sequence, concentration, and transfection time may need optimization to reach satisfactory efficiency.

⁷

CHX is cytotoxic with prolonged incubation time. Working concentration and duration may need optimization to avoid side effects.

⁸

The specificity of primary antibodies is a key to produce high-quality images of immuno-EM, and the most commonly used secondary antibodies are colloidal gold conjugates. Gold particle size varies from 2 to 25 nm.

⁹

The addition of CHX into the release medium can be beneficial to reduce the noise caused by newly synthesized proteins and make it possible to track only the pre-synthesized cargo protein (s).

¹⁰

Prewarming the culture medium to required temperatures is critical for the temperature shift. Water bath is commonly used to ensure a rapid temperature shift and stable temperature maintenance.

¹¹

Both VSVG and RUSH approaches rely on the accumulation of overexpressed cargo molecules in the ER and a wave-like movement of the accumulated molecules along the secretory pathway, and therefore, multiple controls are required to ensure that this overexpression by itself is not changing the trafficking pattern or the properties of secretory organelles.

¹²

The addition of CHX into the release medium is necessary since the analysis depends on the ratio of the two bands. Newly synthesized target protein increases the proportion of the endo H-sensitive form and therefore causes underestimation of the trafficking rate.

¹³

Sucrose concentration is important, so a refractometer is recommended to determine sucrose concentration.

¹⁴

The mitotic index should be examined and a high mitotic index (>90%) ensures the high quality of the mitotic cytosol.

¹⁵

Gene set enrichment analysis (GSEA) uses a walk method in which the score increases for a match between the gene set and ranked input and decreases for no match. This enrichment score is used to determine the leading-edge subsets which represent the most novel matching gene sets. Default ranking is signal to noise for the input dataset. GSEA requires input data normalization, which can be done in DEseq2.

¹⁶

Enrichr uses a simpler matching system than GSEA because it does not consider the rank in the input gene list. It can analyze either crisp datasets, in which genes are either in or not in the gene set, or fuzzy datasets, in which each gene requires manual scoring of the degree of association to the gene set.

¹⁷

Revigo utilizes a semantic similarity algorithm to cluster summarized sets of gene ontologies. Redundant terms are removed or included under parent terms. Summarized lists are then clustered based on ontology or gene set similarity.

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PERMALINK

Common Assays in Mammalian Golgi Studies

Jie Li

Jianchao Zhang

Sarah Bui

Erpan Ahat

Divya Kolli

Whitney Reid

Lijuan Xing

Yanzhuang Wang

Abstract

1. Introduction

2. Materials

2.1. Common Reagents

Table 1. Common reagents used in mammalian Golgi studies.

2.2. Tools Used in RNA-seq Bioinformatic Analysis

2.3. Tools for RNA-seq Analysis and Database Search

2.4. Visualization Tools for RNA-seq Analysis

3. Methods

3.1. Protein Expression, Depletion, and Turnover Assay

3.1.1. Protein Expression by Transient Transfection

3.1.2. Establishment of Stable Cell Lines Using Viral Systems

Table 2. Comparison of non-viral and viral systems in protein expression.

3.1.3. Gene Knockdown

3.1.4. Cycloheximide (CHX) Treatment to Determine Protein Turnover

3.2. Imaging Techniques for Morphological Studies

3.2.1. Immunofluorescence Microscopy

3.2.2. Electron Microscopy (EM)

3.2.3. EM Tomography

3.3. Common Assays to Study Golgi-Mediated Trafficking

3.3.1. Brefeldin A Treatments and Washout

3.3.2. Temperature Block and Release of VSV-G tsO45

3.3.3. Synchronized Release of Cargo Proteins Using the Retention Using Selective Hooks (RUSH) System

3.3.4. Pulse-Chase Labeling and Endo H Treatment

3.4. Subcellular Fractionation and In Vitro Reconstitution Using Isolated Golgi Membranes

3.4.1. Differential Centrifugation to Separate Different Cellular Structure

3.4.2. Equilibrium Gradients to Fractionate Membrane Organelles

3.4.3. Purification of Golgi Membranes from Animal Tissues

3.4.4. In Vitro Budding Assay

3.4.5. In Vitro Reconstitution of Golgi Disassembly and Reassembly During Cell Division

3.4.6. In Vitro Reconstitution of Golgi Structure Formation and Function in Semi-Permeabilized Cells

3.5. Systems Approaches in Golgi Studies

3.5.1. Proteomic and Secretomic Analysis

Fig. 1.

3.5.2. RNA-seq and Transcriptomic Analysis

Fig. 2.

Table 3. Tools and databases for analyzing RNA-seq data.

Table 4. Visualization tools for analyzing RNA-seq data.

3.5.3. Proximity Labeling Using BioID, TurboID, and Apex

Fig. 3.

Acknowledgments

Footnotes

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