Unraveling protein dynamics to understand the brain – the next molecular frontier

Tracers

Protein labeling has a long history with some tools still remaining relevant today. Evans blue (EB) described 100 years ago in 1920 is an example of a dye that is still widely used, in fact even more frequently over the past decade, as it is an inexpensive and clear way to visualize the permeability of organs, critical to understanding the BBB [15, 16]. EB functions by binding albumin, the most abundant protein found in blood plasma. Under normal physiological conditions in young mice, albumin effectively does not cross the BBB. However, damage to the BBB increases the permeability of blood proteins into the brain, which can be visualized by increased EB staining in brain tissue during ischemic stroke, for example [17]. Leakage of intravenously injected EB into the brain was also reported during normal aging in 12 to 24 month old mice, but not 2 to 4 month old mice, suggesting EB is also a useful tool even under more subtle contexts, although care should be taken as old brains often exhibit autofluorescence that may confound the observation of fluorescence from EB [8, 18]. In addition, lung endothelial cells can be used as positive controls for tracers or proteins in BBB experiments as they do not possess the restrictive barrier seen in the brain [19, 20]. Other tracers are available to also track leakage through the BBB, including fluorescein, trypan blue, biotin, horseradish peroxidase, IgG, and isolectin [15, 19]. Labeled dextrans are a tracer of particular interest as the size (10 to 70 kDa), wavelength, and brightness can be easily controlled to examine how different molecular weight molecules interact with the BBB [19, 20].

MRI contrast agents also were originally used in free circulation, especially Gd3+ [21, 22]. Like EB, Gd3+ demonstrates some weak protein binding, often for proteins that naturally coordinate other ions [23]. To avoid accumulation in the body, chelators such as DTPA and DOTA are now often used to facilitate contrast agent removal [24]. These chelators can also be used to directly chemically attach Gd3+ or other lanthanides to a protein of interest to track specific proteins, rather than being a simple free circulating contrast agent [25, 26]. Positron emission tomography (PET) using radionuclides, including 64Cu, 68 Ga, and 177Lu, chelated by DOTA can similarly be used to directly visualize proteins and cells in vivo [27, 28].

Genetically encoded labels

Tools to label proteins have been long used to track individual proteins, of which many are genetically encoded (Fig. 1A). Most commonly, GFP and other fluorescent proteins can be co-expressed attached to a protein of interest as a fusion protein [29]. Although one of the most simple methods of protein labeling, this approach still has advanced uses, such as in super-resolution and single-molecule live-cell microscopy [30, 31]. Intracellular localization of proteins can often be observed via this approach, as well as changes in localization upon various stimuli. In terms of obtaining physiologically accurate results, the major drawback to these fusion proteins is the large size of the fluorescent protein label which can often interfere with biological functions, such as secretion, enzymatic activity, or interactions [32].

Fig. 1figure 1

Tools for protein labeling. Summary of important protein labeling methods discussed in the review and their key advantages and limitations are highlighted here

Alternative labels to GFP fusion proteins include epitope tags such as FLAG, hemagglutinin (HA), and Myc tags [33, 34]. These tags are only a few amino acids, greatly reducing the probability of interfering with protein function. The disadvantage here is that exogenous antibodies must be added for detection, increasing the complexity of the experiment. However, highly specific antibodies for these tags are readily available, minimizing off-target labeling for proteins where antibodies are untested or unavailable and enabling fast and accurate experimental results.

Some peptide tags allow enzymatic modifications to target proteins, such as methods using formylglycine-generating enzyme (FGE), sortase, biotin ligase, and lipoic acid ligase [35]. In these methods, a short peptide sequence is introduced into a specific site in the protein of interest and subsequently acted upon by the enzyme and any additional substrates to generate a new chemical group that can be selectively labeled. For example, FGE recognizes a 6–13 residue peptide tag and converts a cysteine within the motif into an aldehyde group that can be further tagged with probes using hydrazide and aminooxy chemistry [36]. These chemoenzymatic methods are useful for site-specifically installing a number of different functionalities onto proteins but have limited uses in vivo.

Other genetically encoded tags such as glutathione-S-transferase (GST), chitin binding protein (CBP), maltose binding protein (MBP), and His tags should generally be avoided for protein tracking, although they have an advantage in affinity protein purification [33, 34]. GST, CBP, and MBP are all proteins that may interfere with protein function and distribution like GFP, but without the advantage of fluorescence. Also, His tags are small like FLAG, HA, and Myc tags, but, in our experience, antibodies may not always be as effective in specific His tag identification.

SNAP-tags, CLIP-tags, and HaloTags are self-labeling protein tags that provide another alternative to genetically encoded fluorescent proteins [37, 38]. These tags allow for the attachment of a small molecule fluorophore, biotin, or bead to a protein of interest by covalently linking these molecules with the chemical group that reacts with the genetically encoded tag. This application can be useful in specialty applications for some techniques, such as a correlative light and electron microscopy [39]. Of note is that HaloTags have been reported to have ninefold higher signal than SNAP-tags, which can be important for super-resolution microscopy [40].

The major advantage of genetically encoded tags is the high degree of control over the location of the tag within the protein. Small peptide tags have allowed these labels to be applied with minimal interference to protein activity or distribution and to be functionalized with diverse molecules useful in many contexts. While these tools have been successfully used for site-specific labeling of individual proteins, they cannot easily be applied for efficient labeling of many proteins at a time and often face limitations for use in vivo.

Chemically linked labels

Chemical labeling of proteins is a useful way to label specific proteins or even entire proteomes (Fig. 1B). Chemical labeling allows for smaller, potentially less disruptive tags for downstream analysis in experiments [5, 41]. Through chemical labeling, the protein or proteome can be enriched and/or identified at a later time and, in this way, tracked from its original source.

Two of the most widely used chemical bioconjugation methods for proteins are N-hydroxysuccinimidyl (NHS) esters and maleimides [42]. NHS esters label primary amines, targeting lysine side chains and the N-termini of proteins and peptides. However, care should be taken as other residues such as tyrosine and serine can be labeled if reactions occur for too long or at too high concentrations [43]. Maleimides label the side chain of cysteines, but oxidation of cysteines into disulfides can block the reaction [44]. Both reactions can be used independently to confirm that the chemical modification does not interfere with protein function. In our experience, NHS esters are easier to use at first as the reaction is more reliable, especially since lysines often occur on the surface of proteins and cysteines are more likely to be inaccessible [5]. Maleimides can then be used to confirm findings. Maleimides can also be used for site-specific labeling of proteins by first mutating all cysteine residues to serine, which is chemically similar, and then creating cysteine point mutations throughout a specific protein of interest [45]. In addition to lysine and cysteine modifications, many chemical reactions have been developed to specifically label other residues including methionine, tyrosine, aspartate, glutamate, and the N- or C- terminus. Overall, these techniques have been tuned to have increased control on site selectivity but are still limited in terms of specificity of labeling compared to genetically encoded tags.

Critical for visual and spectroscopic tracking of proteins in many applications is the selection of small molecule fluorophores. Small molecule fluorophores may interact with proteins, lipids, and other biomolecules through hydrophobic and electrostatic interactions, leading to inaccurate measurement of fluorescent signals [46]. The brightness of the molecule, protein concentrations, and wavelength, such as autofluorescence of tissues at the GFP/488 nm wavelength, are also important considerations. In our experience, bright Atto dyes in the red shifted region generally give the best results, but controls using other dyes or other proteins should be included to minimize false positives. In addition to fluorophores, many other chemical tags can be linked to proteins using the methods described above, including crosslinkers and enrichment tags. Chemical labeling is very effective for labeling cell surface proteomes and fluid solutions such as blood plasma or CSF that can be labeled ex vivo and re-injected in vivo if needed. Our research group has applied these methods to label the mouse plasma proteome and assess changes in BBB transport during aging in vivo [5].

In vivo SILAC and noncanonical amino acid labels

Methods that minimize changes to proteins are typically less perturbative to protein localization and function. The most non-perturbative labeling method is nonradioactive, isotopic labeling, such as SILAC (Fig. 1C). This method enables whole proteome labeling via the incorporation of amino acids containing heavy isotopes such as 13C, 15 N, and 2H [47]. Heavy-labeling of proteins can then be detected using mass spectrometry based on a defined mass shift compared to the naturally occurring lighter isotopes in proteins. While SILAC is easily carried out in cell culture by growing cells in media containing heavy amino acids, labeling in vivo in mice or other mammals is more difficult due to the expense required to provide several weeks or months of chow for labeling and the inability to easily remove natural isotopes. However, 13C6-Lysine feed has been shown to provide sufficient labeling in mice after about 2 weeks, at a cost for feed of ~ $700 per mouse, which may be a reasonable cost for some experiments [48]. While SILAC is extremely useful for distinguishing proteins from different sources for mass spectrometry applications, there are two main disadvantages of this technology. Firstly, SILAC-labeled proteins cannot be functionalized easily for other applications such as histology. Secondly, and perhaps more importantly, SILAC labeled proteins cannot be enriched, so isotopically-labeled proteins at very low concentrations cannot be easily detected, even with mass spectrometry.

A new alternative to SILAC mice is the use of noncanonical amino acids (ncAAs) which can be incorporated into proteins using native or engineered protein translation machinery (Fig. 1C). ncAAs are often used to introduce bioorthogonal tags that can be labeled or enriched using click chemistry. Azidohomoalanine (AHA) and homopropargylglycine (HPG) are methionine analogs that can be incorporated into wild type mouse or human cells and tissues by endogenous methionyl-tRNA synthetases, with minimal changes to protein biology [49, 50]. AHA or HPG can be provided as feed, in drinking water, or injected, but we have noticed toxicity with intraperitoneal injections in mice, especially at concentrations at the maximum levels of what was reported previously [50]. These ncAAs are best used when provided in mouse feed containing 0% methionine to reduce methionine incorporation [49]. We have successfully used both AHA and HPG to label tissues and blood plasma, and, in our experience, AHA labeling is noticeably stronger and less toxic than HPG, so AHA is greatly preferred. One disadvantage of AHA/HPG mouse labeling is that specific cell types or tissues cannot be targeted as these ncAAs use the native mouse methionyl-tRNA synthetase for incorporation, so labeling occurs throughout the entire mouse. We also observe moderately less labeling in the brain, likely due to the BBB occluding the ncAAs.

To overcome the challenge of tissue-specific or cell-specific incorporation of ncAAs in animals, the use of the pyrrolysyl-tRNA synthetase/tRNA pair (PylRS/tRNAPyl) that targets the amber codon was first implemented, as only cells or tissues expressing this orthogonal pair will be labeled with the corresponding ncAA [51]. Since PylRS/tRNAPyl does not occur in mammals, it can incorporate novel ncAAs into cells or tissues only where it is expressed. PylRS/tRNAPyl has mostly been used to introduce ncAAs site-specifically into proteins genetically modified to contain the amber codon in E. coli until recently, but this system has an advantage of now > 150 ncAAs engineered for use through PylRS mutations [51, 52]. The lab of Jason Chin has expanded the PylRS/tRNAPyl pair into mice and other eukaryotes, with their SORT-M system. This technology can also target any amino acid codon, rather than only the amber codon, by mutating the tRNAPyl anticodon to correspond to the targeted codon. This system has also been further improved by mutations that increase the tRNAPyl stability in mammalian cells [52]. Two versions of ncAAs, one containing a methylcyclopropene side chain and one containing a terminal alkyne can be used with the SORT-M system to facilitate bioorthogonal click chemistry with desired probes. The SORT-M system is the most diverse system for use in eukaryotes, although in our hands we have found it to be less robust for strong labeling as the tRNA stability is still limited in vivo. However, this system has now been clearly demonstrated to be a powerful tool in mice with recent method developments of the Chin lab [53].

To overcome the limitation of tRNAPyl stability or expression, certain aminoacyl-tRNA synthetase (aaRS) mutants exist that can incorporate ncAAs into mammalian cells without requiring the expression an exogenous tRNA. The lab of David Tirrell found a mutant mouse methionyl-tRNA synthetase (MmMetRSL274G), allowing the incorporation of the azide-containing azidonorleucine (Anl) at methionine sites in mice [54, 55]. We have also recently discovered a mutant yeast tyrosyl (ScTyrRSY43G) and mouse phenylalanyl (MmPheRST413G) tRNA synthetase that can incorporate 3-azidotyrosine at tyrosine sites and 4-azidophenylalanine at phenylalanine sites, respectively, in mice [56, 57]. The MmMetRSL274G, ScTyrY43G, and MmPheT413G all are efficient at amino acid incorporation in our hands. However, we have observed the MetRS and TyrRS mutants seem to have the most robust incorporations of their respective ncAAs in mice compared to PheRS or the SORT-M system. The TyrRS also has the advantage of a short gene sequence, which is important for the limited packing capacity of adeno-associated viruses (AAVs) for viral genetic transduction. The TyrRS also allows ncAA incorporation at near maximum efficiency even at low concentrations (15 µM of 3-azidotyrosine) [56]. Overall, SORT-M, MmMetRSL274G, ScTyrY43G, and MmPheT413G all have shown great promise for cell and tissue-specific proteome labeling. Using a multiple of these four approaches can also be useful to increase proteome coverage and to ensure reproducibility of results with different ncAAs incorporated into different amino acid sites.

The lab of Alice Ting has also produced other complementary methods using proximity labeling with BioID or TurboID, which both utilize a biotin ligase linked to a protein of interest or cellular localization sequence to attach biotin to nearby surrounding proteins (Fig. 1D) [58,59,60]. As pulling down biotinylated proteins using streptavidin is a widely-used, low background, and high-specificity technique, it can offer easy adoption and reliable results. One caveat is that endogenous biotin is consumed by the method, so care must be taken especially when used in vertebrates. There are also endogenously biotinylated proteins, but these can be subtracted out as they are also present in control samples. This approach has been used in vivo to successfully label tissue-specific secretomes in mice, which can then be detected in serum [60]. This approach also allows for a powerful proximity labeling technique, in which the biotin ligase is attached to a single protein to identify the nearby interaction partners, even revealing many interaction partners that are unknown [61].

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