Highly Sensitive and Multiplexed Protein Measurements - Chemical

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Cite This: Chem. Rev. XXXX, XXX, XXX−XXX

Highly Sensitive and Multiplexed Protein Measurements Limor Cohen†,‡,§ and David R. Walt*,†,‡ †

Department of Pathology, Brigham and Women’s Hospital, Harvard Medical School, Boston, Massachusetts 02115, United States Wyss Institute for Biologically Inspired Engineering and §Department of Chemical Biology, Harvard University, Boston, Massachusetts 02115, United States

Chem. Rev. Downloaded from pubs.acs.org by UNIV OF SOUTH DAKOTA on 08/29/18. For personal use only.

‡

ABSTRACT: Proteins are involved in many biological processes. Misfolded, truncated, or mutated proteins as well as over- or underexpressed proteins have been implicated in many diseases. Therefore, detection and quantification of proteins is extremely important. Conventional techniques such as the enzyme-linked immunosorbent assay, Western Blot, and mass spectrometry have enabled discovery and study of proteins in biological samples. However, many important proteins are present at low concentrations, rendering them undetectable using conventional techniques. Furthermore, limited ability to simultaneously measure multiple proteins in a sample has constrained our ability to fully study the proteome. In this review, we comprehensively discuss approaches for protein detection. We first discuss the fundamentals of proteins and protein assays, including affinity reagents, surface functionalization, assay formats, signal detection, and multiplexing. We then discuss the challenges with these methods and review existing methods for highly sensitive and multiplexed protein detection. Finally, we review recent advances in protein detection from the literature and discuss challenges and future directions.

CONTENTS 1. Introduction 2. Proteins 3. Fundamentals of Protein Assays 3.1. Affinity Reagents 3.1.1. Antibodies 3.1.2. Aptamers 3.1.3. Other Affinity Reagents 3.2. Surface Functionalization 3.3. Assay Formats 3.4. Signal Detection 3.5. Multiplexing 4. Current Commonly Used Methods for Protein Detection 4.1. Detection of Total Protein 4.2. Enzyme-Linked Immunosorbent Assay (ELISA) 4.3. Enzyme-Linked Immunospot (ELISPOT) 4.4. Western Blot 4.5. Protein Microarrays 4.6. Flow Cytometry 4.7. Proximity Ligation Assay (PLA) and other Nucleic Acid-Based Detection Methods 4.8. Mass Spectrometry 4.9. Lateral Flow Assay (LFA) 4.10. Surface Plasmon Resonance (SPR) 4.11. Optical Imaging 5. Challenges with Protein Detection 6. Emerging Methods for Ultrasensitive and Highly Multiplexed Measurements 6.1. Ultrasensitive Methods for Protein Detection © XXXX American Chemical Society

6.1.1. Meso Scale Discovery (MSD) 6.1.2. Single-Molecule Arrays (Simoa) 6.1.3. Single-Molecule Counting (SMC) 6.1.4. Single-Cell Western Blot 6.2. Highly Multiplexed Methods for Protein Detection 6.2.1. SOMAscan Assay 6.2.2. Luminex 6.2.3. Mass Cytometry (CyTOF) 7. Advances from the Literature 7.1. Advances in Labeling and Signal Detection 7.1.1. Upconverting Nanoparticles 7.1.2. Photoelectrochemical Detection (PEC) 7.1.3. Optical Ring Resonators 7.1.4. Surface-Enhanced Raman Scattering (SERS) Tags 7.2. Miniaturization 7.2.1. Microcantilever-Based Assays 7.2.2. Ultrasmall Containers 8. Conclusion and Future Directions Author Information Corresponding Author ORCID Notes Biographies References

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Special Issue: Chemical Sensors

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Received: April 20, 2018

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DOI: 10.1021/acs.chemrev.8b00257 Chem. Rev. XXXX, XXX, XXX−XXX

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1. INTRODUCTION Proteins are macromolecules involved in important biological functions including replicating genomic information, regulating transcription, signaling, providing structure, catalyzing reactions, and transporting molecules. Many proteins are dysregulated and differentially expressed in disease and can serve as drug targets or biomarkers. Therefore, tools for measuring and detecting proteins are essential for fully characterizing these functions. Several challenges exist with protein detection methods. First, a sample may contain many interfering molecules that make it difficult to detect a specific protein. Second, many proteins may be present at low concentrations in a biological sample. Third, concentrations of different proteins can vary by many orders of magnitude. Finally, protein processing, such as post-translational modifications and spliced variants that result in different isoforms, can make it difficult or complicated to detect the specific protein of interest. Common techniques currently employed for protein detection include the Western blot, mass spectrometry, and the enzyme-linked immunosorbent assay (ELISA). These techniques have provided a wealth of information on proteins and their function; however, they may suffer from shortcomings in throughput, multiplexing capabilities, specificity, and sensitivity. In the past few years, many tools have been developed to overcome these challenges. In this review, we comprehensively discuss existing and innovative analytical tools for protein detection and quantification. First, we briefly discuss the fundamental properties of proteins. We then discuss fundamentals of protein assays, including affinity reagents, surface functionalization, assay formats, signal detection, and multiplexing. We then review commonly used methods for protein detection. We then discuss the challenges with these methods and review existing methods for highly sensitive and multiplexed protein detection. Finally, we review advances in protein detection techniques from the literature and discuss challenges and future directions.

ubiquination, deamidation, oxidation, sulfation, and nitration. The five major classes of covalent modifications are shown in Figure 1.4 In some cases, detection of a specific post-

Figure 1. Five major types of covalent modifications to protein side chains: phosphorylation, acylation, alkylation, glycosylation, and oxidation. Reprinted with permission from ref 4. Copyright 2005 John Wiley and Sons.

translational modification on a protein may be desirable over detection of the unmodified protein because it may be the functional or disease-causing species. Some protein detection methods first start by treating the protein with a glycosylase, which removes glycans. This treatment is designed to reduce the complexity of the sample but may remove important PTMs that are key to understanding the sample being measured. These modifications play critical roles in the function of proteins and can be dysregulated in disease.5−9 For example, approximately 30% of proteins encoded by the human genome are phosphorylated, with irregular phosphorylation patterns associated with many human diseases.10 Thus, it is not only important to detect specific proteins but also to detect specific modifications on certain proteins. Additionally, proteins range in size from very small monomeric proteins to large macromolecular complexes, in which proteins interact with other molecules such as RNA, DNA, or other proteins. Therefore, it is often challenging to detect a specific protein or a specific modification on a protein in a complex biological sample. Second, some proteins of interest may be present at low levels, limiting the ability to detect them. It can be particularly difficult to measure low levels of a given protein in a complex biological sample that contains high levels of other potentially interfering molecules. Third, it is often necessary to detect more than one protein in a single sample. Many proteins work in networks to exert their function. Some proteins are

2. PROTEINS Proteins are a class of biological macromolecules that play important roles in many biological processes. Proteins are essential for replicating genomic information, regulating gene expression, transcribing mRNA, signaling, catalyzing metabolic reactions, and transporting molecules. Disrupted protein expression as well as dysfunctional mutated or misfolded proteins are associated with multiple diseases, such as Alzheimer’s disease and sickle cell anemia. Thus, detecting specific proteins is important. Several features of proteins are important for their detection and are discussed in this section. First, the proteome consists of tens of thousands of proteins, and thus, detecting a specific protein in a complex matrix is challenging. The Human Genome Project revealed that different proteins can be encoded by the same genes. It has been estimated that the human genome contains approximately 20 000 protein coding genes,1 yet the number of proteins is considerably higher as a result of variations in DNA sequence, alternative mRNA splicing, post-translational modifications, and proteolytic cleavage.2 A mammalian cell contains a total of 109 protein molecules, and a bacterial cell contains a total of 106 protein molecules.3 Proteins are modified by covalent post-translational modifications including phosphorylation, acetylation, methylation, acylation, glycosylation, B


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3.1.1. Antibodies. Antibodies, major components of the vertebrate immune system, bind to foreign molecules known as antigens. Most antigens are proteins. An antibody binds specifically to a structure on the antigen called an epitope. Several classes of antibodies exist, with Immunoglobulin G (IgG) being a major class and most commonly used in bioanalytical applications. IgGs have four polypeptide chains consisting of two heavy chains and two light chains that form a 150 kDa molecular complex (Figure 2). The variable domains

overexpressed, while others are under expressed. Protein measurements on the omics scale are highly advantageous and provide unbiased detection of proteins. Measurements of many proteins simultaneously is challenging because proteins in a biological sample can differ in concentration by several orders of magnitude.11 Additionally, levels of a target protein can vary substantially between different biological samples, which further complicates detection. Finally, the particular structural form of a protein may depend on its native cellular context. To analyze proteins, it is often necessary to alter their native environment or remove them from it completely. For example, hydrophobic membrane proteins that are embedded in the lipid bilayer may not be stable in aqueous environments. Therefore, when removing a protein from its native environment for analysis, it is important to determine if such manipulations will affect the analysis.

3. FUNDAMENTALS OF PROTEIN ASSAYS Methods to analyze proteins can be broadly classified into three categories. The first category is protein detection using solution-based assays, which will be the focus of this review. The second category is protein detection using mass spectrometry-based methods. The third category is structural analysis of proteins using methods such as X-ray crystallography, NMR, Cryo-EM, and circular dichroism (CD). The latter two categories have been thoroughly reviewed elsewhere and will not be the focus of this review.12,13 A protein assay typically consists of three major components: an affinity reagent, a signal transducer, and a detector.14 The role of the affinity reagent is to specifically bind to the target protein molecule. This binding enables significant purification of the protein from the complex sample being analyzed. The signal transducer then converts the binding event into a measurable signal, such as an optical, electrochemical, or mechanical signal. The detector, such as a chargecoupled device (CCD), then converts the signal into a digital readout. In this section we discuss affinity reagents, surface functionalization, assay formats, signal detection, and multiplexing.

Figure 2. IgG structure showing two different molecular depictions on the left and right sides. IgGs consist of two heavy chains and two light chains that are linked by a disulfide bond. IgG contains two antigen-binding sites. Modified and reprinted with permission from ref 20. Copyright 2017 Annual Reviews.

of the light and heavy chains associate to form the antigenbinding domain, also known as the Fab. In 1993, it was discovered that sharks16 and camelids, such as camels and llamas,17 produce antibodies that do not have a light chain. These single-domain antibodies are known as Nanobodies or VHHs (Figure 3).18 Single-domain antibodies have lower molecular weights (90 kDa) than traditional antibodies (150 kDa) and therefore may be better affinity reagents.19 Antibodies are generally produced by affinity maturation in a living organism. Antibody diversity is attained by combinatorial biosynthesis based on genetic recombination capable of

3.1. Affinity Reagents

An affinity reagent is a molecule that recognizes a specific protein. Binding of the protein to the affinity reagent is converted into a measurable signal. Affinity reagents may need to be adsorbed or covalently attached to a surface. Approaches for covalent conjugation of biomolecules have been thoroughly described elsewehere.15 Additionally, affinity reagents need to interact with the signal transducer either directly or via a secondary molecule that can interact with the signal transducer. These interactions must not affect the performance of the measurement by sterically hindering binding to the protein or affecting the performance of the signal transducer. There are several important characteristics of protein affinity reagents including high affinity (low dissociation constants), high specificity (ability to recognize the target protein in a sample containing many other potentially interfering molecules), cost, stability, and reproducibility. The most commonly used affinity reagents for proteins are antibodies and aptamers. Other affinity reagents such as molecularly imprinted polymers, lectins, peptides, and antibody mimetics can also be used. These affinity reagents will be discussed in this section.

Figure 3. Single-domain antibody structure. Single-domain antibodies are much smaller compared to IgGs. Single-domain antibodies do not have a light chain and are derived only from the heavy chain. Singledomain antibodies are naturally found in camelids and sharks. (v, variable; HCAb, heavy-chain-only antibodies; H, heavy chain). Modified and reprinted with permission from ref 21. Copyright 2014 Nature Publishing Group. C


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producing more than 108 different antibodies that have different binding specificities.22 The most common antibody preparations are polyclonal and monoclonal antibodies. Polyclonal antibodies are generated in vivo via affinity maturation. Therefore, polyclonal antibodies recognize different epitopes of a given antigen. Monoclonal antibodies are derived from a hybridoma, a fusion of a B cell with an immortal cell line, and were first described in 1975.23 Since monoclonal antibodies are derived from a clone of identical B cells, they recognize a single epitope. In some cases, it is preferable to use one clone over another; for example, when detection of a specific post-translational modification is required, a monoclonal preparation is necessary since a polyclonal preparation will detect all of the epitopes on the protein. On the other hand, in some cases a polyclonal preparation is favorable−for instance, when detecting the presence of all proteins containing a particular epitope, regardless of the PTM. A disadvantage of polyclonal antibodies is the potential lack of reproducibility between different reagent lots. Once a polyclonal preparation is depleted an animal must be immunized again to produce more polyclonal antibodies. The second polyclonal batch will likely have a different composition and may perform differently than the first batch, requiring new assay conditions to be determined and new calibrations to be performed. Several approaches for in vitro antibody production have been developed. These include phage, yeast,24 and ribosome display,25,26 which have been reviewed elsewhere.24,27,28 These approaches are advantageous for high-throughput antibody production as well as production of antibodies against toxic molecules and molecules that do not elicit an immune response in an animal. Another advantage is the ability to incorporate a tag, such as a his-tag, for antibody immobilization and conjugation to various labels. Finally, these recombinant preparations may reduce the cost of antibody production. Antibody−antigen interactions are particularly strong and have low KD values. In vivo affinity maturation, with repeated exposure of an animal to the same antigen, typically results in antibodies with affinities in the nanomolar to picomolar range,29 but directed evolution of antibodies with affinities in the femtomolar range have been reported.30 Antibodies are generally highly specific to their target, although nonspecific binding can occur. Overall, due to their high affinity and specificity, antibodies are key reagents for protein detection.31 Antibodies can be easily integrated into protein assays. Antibodies have multiple reactive side chains that can serve as attachment sites for conjugation to solid supports, such as magnetic beads or nanoparticles, or labels that act as signal transducers. Antibodies are usually covalently attached to surfaces or detection labels. Another approach is to utilize biotinylated antibodies that can interact with streptavidin, which has been attached to a surface or a signal transducer. However, covalent conjugation may affect interaction with the target protein. Modifying a molecule that can interact with the antibody may overcome this challenge. A secondary antibody, Protein A, or Protein G, which bind to the Fc region of the antibody, may be used. 3.1.2. Aptamers. Aptamers are single-stranded DNA or RNA oligonucleotides that can bind to proteins with high affinity and specificity.32,33 First described in 1990,34,35 aptamers are created by an in vitro process known as SELEX, systemic evolution of ligands by exponential enrich-

ment. The SELEX process starts with a large combinatorial library of DNA or RNA oligonucleotides of random sequences. These sequences are added to the target protein, usually attached to a solid support, and some bind with high affinity. The sequences that do not bind to the target protein are then separated from those that bind to the target protein. The bound sequences are eluted from the protein, amplified, and undergo another round of selection. This process is repeated, typically between 8 and 15 rounds, with each iteration subjecting the bound sequences to higher stringency to elicit stronger binders. The targets are then cloned and sequenced. Aptamers have several advantages for use as affinity reagents such as low cost, high stability, and high specificity, with dissociation constants in the femtomolar to picomolar range.36−38 Once an optimal sequence has been determined, they can be easily and reproducibly generated by chemical synthesis. Additionally, DNA aptamers are highly stable reagents such that bound proteins can be removed either chemically or thermally to regenerate the free aptamer for another round of binding. Another advantage of aptamers is that they can be generated against almost any protein, even toxic proteins and those that are not immunogenic. Aptamers can be easily integrated into many protein assay formats by substituting for antibodies and are easily labeled and modified with various molecules and functional groups. Aptamers are highly stable and can also be regenerated for numerous uses. The SOMAscan assay, which is described in section 6.1.1, uses aptamers as affinity reagents. 3.1.3. Other Affinity Reagents. Other than antibodies and aptamers, additional affinity reagents are available including molecularly imprinted polymers (MIPs),39,40 protein ligands, lectins, and antibody mimetics. The protein itself or other peptide ligands can be used to detect protein−protein interactions. For example, when one needs to detect antibodies to determine if an immune response has occurred, the protein can be used to bind to the target antibodies. Lectins, such as wheat germ agglutinin or concanavalin A,41 bind to glycans on glycoproteins and are commonly used. Alternative molecules that mimic antibodies, known as antibody mimetics, are proteins that are structurally unrelated to antibodies and can bind to targets specifically through protein-engineering approaches.20 3.2. Surface Functionalization

In many protein assays, the target protein must first come into contact with an affinity reagent that is immobilized on the surface. Therefore, the performance of a protein assay is highly dependent on the chemical modification of the surface with an affinity reagent.42 Approaches to attach an affinity reagent onto the surface include noncovalent and covalent immobilization. Strategies for covalent immobilization chemistries have been thoroughly described.15 Noncovalent surface attachment can be achieved by direct electrostatic interactions between the surface and the affinity reagent. Alternatively, secondary molecules such as biotin−streptavidin or attachment via interaction of Proteins A and G to the Fc region of the antibody can also be used. Self-assembled monolayers (SAMs) are also commonly used to attach affinity reagents to a surface.43 Finally, encapsulation of the affinity reagent is another approach. All of these strategies can be used to attach an affinity reagent to the surface (Figure 4).44 There are two different implementations for using surfacebound affinity reagents. The first implementation is one in D


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which the target protein must diffuse to the surface, for example, when using an affinity reagent immobilized onto the surface of a traditional microtiter plate. The second implementation is when the affinity reagent is immobilized onto the surface of micro- or nanoparticles and the target protein and the surface are both in solution. Solution-based assays enhance and accelerate the binding kinetics and capture efficiency of the target protein due to superior mass transport of the protein to the surface. Another advantage is the higher surface to volume ratio of micro- and nanoparticles, which further improves the capture efficiency. Another consideration is the orientation of the affinity reagent on the surface and the surface density or number of affinity reagents per area. Both have been shown to affect the binding efficacy of the target protein.46,47 Recently, several studies have assessed the relationship between capture efficiency and antibody functionalization of gold nanoparticles.45,48−51 Due to their favorable properties, nanomaterials are becoming more popular for use in protein detection techniques. These nanomaterials include carbon nanotubes, polymer nanowires, and zinc oxide nanorods.52−55 Reduction of nonspecific binding to the surface is also important. Methods for passivating the surface include using BSA56 or various neutral or hydrophilic macromolecules such as poly(hydroxyethyl methacrylate), poly(acrylamide), and poly(ethylene glycol).57−60 Finally, another consideration is the ability to regenerate the binding capabilities of the surface for multiple independent measurements of proteins.61 Regeneration is achieved by overcoming the attractive forces between the affinity reagent and the bound protein, ideally without destroying the binding capabilities of the former. Strategies to regenerate the binding

Figure 4. Five strategies used for bioconjugation of affinity reagents to surfaces via (i) electrostatic interactions, (ii) direct interaction with the surface, (iii) secondary interactions, (iv) covalent immobilization, and (v) encapsulation. Reprinted with permission from ref 45. Copyright 2013 American Chemical Society.

Figure 5. Immunoassay formats. (a) Label-free immunoassay. Binding of the target protein to capture antibodies immobilized onto a surface results in a measurable signal. (b) Sandwich immunoassay in which the detection antibody is labeled directly with a tag such as a fluorophore. (c) Sandwich immunoassay in which an enzyme is used to amplify the signal. (d) Competitive immunoassay in which the target protein competes with the labeled protein from binding to the capture reagent. (a−d) Heterogeneous immunoassays in which the capture reagent is immobilized onto a surface and the unbound sample components are washed away. (e) Homogeneous immunoassay in which the reagents are in solution and are not separated from other unbound sample components. E


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capabilities of surfaces have been developed;62 however, it is still quite challenging to regenerate the surface because the performance is compromised due to reduced binding affinity, increased nonspecific binding, and thus lower sensitivity. Some affinity reagents, such as aptamers, may be more amenable to regeneration than antibodies. The main benefits of surface regeneration are reduced cost and the ability to monitor changes in concentration over time without changing the affinity capture surface.

Optical detection is most commonly used for signal transduction. Optical detection is a signal transduction method based on measuring changes in the amount of light absorbed or emitted at a specific wavelength, typically in the ultraviolet (UV) or infrared (IR) regions. The affinity reagent is labeled with a molecule that can be optically detected. Since the ability to detect the protein analyte depends on detecting the optical label, the label needs to either have a high extinction coefficient for absorbance measurements or have a high quantum yield for fluorescence emission measurements. In addition, there should be a sufficient degree of labeling on the affinity reagent. Fluorescent labels are more commonly used in optical detection. Two key properties of fluorophores are their Stokes shift and quantum yield. A larger Stokes shift results in better signal separation from the excitation wavelength. The quantum yield, along with the molar extinction coefficient, determines the overall brightness of the fluorophore.68 Numerous fluorescent dyes and their derivatives have been developed to increase the brightness and quantum yield.69 The most common fluorescent core structures are water-soluble conjugated organic molecules that are derivatives of cyanine, fluorescein, rhodamine, and coumarin. Derivatives with aromatic ring constituents, such as electron-donating or -withdrawing groups, can have drastic effects on fluorescence and quantum yield. Fluorescent molecules other than small organic dyes can also be used for optical detection. These include naturally fluorescent protein molecules, such as phycoerythrin and green fluorescent protein (GFP). However, these molecules are larger than organic dyes and may sterically hinder the protein from binding to the affinity reagent. There are several limitations to detecting a fluorophore that has been directly conjugated to an affinity reagent. First, the protein-binding site on the affinity reagent may be unavailable for binding to the protein. Second, when multiple fluorophores are proximal to each other, quenching could occur. Finally, a limited number of fluorophores can be attached to an affinity reagent either because more labels will affect the binding affinity or because there are a limited number of sites where the label can react. Typically, only 8−10 fluorophores can be conjugated to an antibody. In general, amplifying the signal leads to enhanced sensitivity. Various approaches to signal amplification have been implemented. One approach is by using an enzyme that generates many detectable signal molecules. Enzymatic turnover amplifies the signal and increases the sensitivity because the number of detectable molecules is substantially higher than the number of protein molecules captured. Commonly used enzymes include alkaline phosphatase (AP), horseradish peroxidase (HRP), and β-galactosidase. These enzymes turn over many substrate molecules to generate many fluorescent molecules. A second approach is to use nanomaterials.70−72 Nanomaterials have increased surface area such that they can be conjugated to both an affinity reagent and many labels, such as fluorophores or enzymes. Nanomaterials that are inherently brighter, such as carbon dots73−75 or polymer dots,76−78 can also be used as optical labels. Fluorescent silica nanoparticles in which standard organic dyes are incorporated can also be used to enhance the signal.79−81 Fluorescent molecules are sensitive to changes in the chemical environment. Buffer composition, pH, and solvent polarity can affect the quantum yield. Overlabeling an affinity reagent with a fluorescent molecule can also change the quantum yield due to quenching resulting from interactions

3.3. Assay Formats

The two most common formats for protein assays are noncompetitive and competitive assays. Noncompetitive assays can be further broken down into label-free assays (Figure 5a) and sandwich assays (Figure 5a−c). Both label-free and sandwich assays require a capture reagent that specifically binds to the target protein. In a label-free assay, binding of the target protein to the capture reagent results in a measurable change. In a sandwich assay, a second affinity reagent binds to a different epitope on the target protein, forming a “sandwich.” The second affinity reagent is labeled with a tag, such as a fluorophore (Figure 5b), or an enzyme (Figure 5c) that produces molecules that can be detected. In a competitive assay (Figure 5d), the surface is coated with a capture reagent that binds to the target protein. A sample containing the target protein and a known amount of labeled target protein are added. The target protein competes with the labeled protein from binding to the capture reagent. When low levels of the target proteins are present relative to the labeled protein, the labeled protein can bind to the capture reagents to produce a signal. With increasing levels of the target protein, binding of the labeled protein to the capture reagent decreases. Therefore, the concentration of the target protein is inversely proportional to the signal. A competitive assay for protein detection is less commonly used than the noncompetitive assay. However, it is often used to detect antibodies or when only one affinity reagent is available. Protein assays can be homogeneous (Figure 5e) or heterogeneous (Figures 5a−d). In a homogeneous assay, the reagents are in solution and do not require separation from other unbound sample components, while in a heterogeneous assay, the capture reagent is immobilized onto a surface and the unbound sample components are washed away. Heterogeneous assays are more commonly used than homogeneous assays. 3.4. Signal Detection

Interaction of a protein with an affinity reagent must be converted into a measurable signal. Signal transduction approaches fall into two general categories: chemical and physical (or label-free) transduction. Chemical transduction is a change in chemical composition coupled to a binding event, for example, when an enzyme catalyzes the formation of a fluorescent product. Physical transduction is a measurable change in the physical properties in response to the interaction of the affinity reagent with the protein. Because physical transduction methods do not require a chemical label, they are also known as label-free transduction. Several reviews have been written that provide more detail on optical label-free detection methods including theoretical discusions.63−67 Chemical and label-free signal transduction approaches relevant to protein detection include optical, electrochemical, and mechanical signal transduction. F


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between the dyes. Additionally, nonspecific binding can arise between organic dyes containing aromatic groups and other components in the assay such as surfaces.82 Such interactions can be minimized by adding negative charges to the fluorophore, such as PEG or sulfonate modifications, to increase the hydrophilicity.15 Naturally occurring fluorophores in a biological sample, such as aromatic amino acid residues, NADH, and flavins, can also cause interference and increase the background. Other commonly used methods for optical detection, particularly as part of the enzyme-linked immunosorbent assay (ELISA), include electrochemiluminescence (ECL), chemiluminescence, and colorimetric-based assays. In ECL, an electron transfer reaction at the surface of an electrode produces excited states that can then emit light. The MSD assay uses ECL for protein detection and is discussed in section 6.1.1. Chemiluminescence is a result of a chemical reaction that produces light. In a chemiluminescent reaction, oxidation of an organic dye, such as luminol, with a strong oxidant, such as hydrogen peroxide, in the presence of a catalyst produces a light-emitting molecule. The emission lifetime of the molecule can last from several seconds to hours. Commonly used chemiluminescent reagents for protein detection are the enzyme horseradish peroxidase and luminol or its derivatives. Chemiluminescent detection does not require an excitation source, and therefore, there is no background signal due to excitation of other components in the sample matrix. A disadvantage is that the generated chemiluminescent signal can be relatively weak; therefore, approaches to enhance chemiluminescent intensity have been developed.83 These approaches include modification of luminol or addition of various organic compounds, such as phenols, that enhance chemiluminescence. Quantum dots and metallic nanoparticles have also been used to enhance chemiluminescence.84−86 For example, it has been shown that gold nanoparticles can catalyze chemiluminescence of luminol.87 Additionally, carbon dots have been shown to have chemiluminescent properties.88 Colorimetric signal generation is based on formation of a colored precipitate, typically by an enzyme.89 Nanoparticlebased nonenzymatic colorimetric detection has also been demonstrated.90,91 Nanoparticle-based colorimetric assays can exhibit high sensitivities for protein markers,92−95 down to the attomolar range.96 Advantages of colorimetric assays include relative simplicity, low cost, and ability to detect markers with the naked eye.97 In addition to an optical label, optical detection tools often require components such as a light source, optical filters, and detectors, such as photodiodes, phototransistors, or photomultiplier tubes, which convert the incident light into an electrical signal. Additional information on instrumentation for optical detection can be found in the literature.68 Electrochemical methods for protein detection are widely used in the literature and have been recently reviewed.98−100

Figure 6. Approaches for multiplexing via (a) a conventional microarray in which the identity of the target protein is known based on the predetermined position of the affinity reagent and (b) a suspension array in which the affinity reagents are attached to encoded particles. Each optically distinct population of beads is conjugated to capture antibodies to a specific protein. Printed with permission from ref 108. Copyright 2006 John Wiley and Sons.

example, the SOMAscan assay, which is described in more detail in section 6.2.1, can detect approximately 1300 different proteins simultaneously using a nucleic acid microarray. The second format employs encoded micro- or nanoparticles, in which each uniquely encoded particle population is functionalized with an affinity reagent specific to a given protein. There are several ways to encode micro- and nanoparticles, including optical encoding with fluorescent dyes or Raman tags.103−105 Luminex has developed a 100-plex assay by optically encoding microspheres with combinations of fluorescent dyes.106,107 Other ways to encode are based on graphical, physical, magnetic, and thermal properties. These methods have been recently reviewed.108−110 Finally, encoding using nucleic acid barcodes can provide high multiplexing capabilities.111 A major challenge with multiplexed protein detection assays is nonspecific binding and cross-reactivity between affinity reagents, secondary labels, and other components in the biological sample.112 Despite many efforts, measuring multiple protein analytes with high sensitivity, specificity, and accuracy is still a major challenge.

4. CURRENT COMMONLY USED METHODS FOR PROTEIN DETECTION This section discusses the most widely used techniques for protein measurements, particularly for life sciences research and clinical diagnostics. Many of these techniques were developed in the 1960s and 1970s and are still widely used today. These techniques include direct detection of total protein, the enzyme-linked immunosorbent assay (ELISA), enzyme-linked immunospot (ELISPOT), Western blot, protein microarrays, flow cytometry, proximity ligation assays (PLAs) and other nucleic acid-based methods, mass spectrometry, lateral flow assays, surface plasmon resonance (SPR), and optical imaging. Improvements in protein detection methods are commonly based on improving these basic techniques. Other commonly used techniques, such as Xray crystallography, CD, NMR, and electron microscopy, will not be covered. These techniques are used to analyze the structure of a specific protein or interactions between two proteins and require purified protein samples. This review focuses on measuring unknown levels of proteins in a biological sample. Several reviews have been recently written on various aspects of protein detection techniques.113−120

3.5. Multiplexing

Measuring multiple different proteins in a single sample simultaneously is advantageous for high-throughput analysis as well as reduced time and cost. There are two commonly used multiplex assay formats (Figure 6). The first format spatially separates the capture reagents using a conventional microarray, in which the identity of the capture reagent at each position in the array is predetermined by its spatial location.101,102 For

4.1. Detection of Total Protein

Direct detection and quantification of total protein is important for many applications. Methods using UV and visible spectroscopy are commonly used for fast quantification G


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methods are advantageous for direct and simple detection and quantification of total protein and are widely in use.

of total protein concentration relative to a standard or based on known protein extinction coefficients. The aromatic amino acids, tryptophan and tyrosine and to a lesser extent phenylalanine, absorb light at 280 nm, a property which enables quantification based on absorbance. This technique is easy to use, but disadvantages include interference from other sample components, such as nucleic acids, and low sensitivity due to low extinction coefficients of amino acid residues. Additionally, different proteins have different absorbance spectra depending on the structure and the number of aromatic amino acid residues, which can complicate quantification and reduce accuracy. Another strategy for total protein quantification is based on colorimetric protein assays. There are two general approaches. The first approach is based on chelation of copper by proteins and includes the biuret assay, the bicinchoninic acid assay (BCA assay), and the Lowry protein assay. In the biuret assay, peptides with three or more amino acid residues form a colored chelated complex with Cu2+ under alkaline conditions. The color intensity is proportional to the number of amino acid residues. The biuret test is a simple and fast quantitative test to measure total protein concentration, with low sensitivities in the mg/mL range. The BCA assay, developed in 1985, can be used to enhance sensitivity.121 It is based on the biuret reaction, in which proteins first form a chelated complex with Cu2+. Cu2+ is then reduced and chelated with BCA to form a colored complex. Another widely used technique is the Lowry test, which was developed in 1951. The Lowry test is also based on the biuret reaction and involves the reduction of copper (Cu2+ to Cu+) by proteins in alkaline solutions and formation of a colored compound via reduction of a color-enhancing Folin phenol reagent.122 The colored compound is produced and can be measured at a long wavelength (750 nm), which is advantageous since fewer compounds absorb light at that wavelength and therefore the background is lower. The second approach is based on protein-binding dyes. The most commonly used assay is the Bradford assay, developed in 1976, which utilizes the dye Coomassie G-250.123 The Coomassie dye binds selectively to certain amino acids and tertiary protein structures. Therefore, there can be variations between dye-binding efficiency to different proteins, which can reduce accuracy. For the dye to bind to proteins, the protein mass must be high, and therefore, this method is generally used for proteins with high molecular weights. The Bradford assay is rapid, simple, and widely used, but its accuracy is limited. Fluorescent dyes can also be used due to their favorable properties including increased sensitivity, decreased background, and wider dynamic range. In this method, a nonfluorescent dye interacts with proteins either covalently or noncovalently. This interaction causes the dye to become fluorescent. 1 2 4 For example, the CBQCA (3-(4carboxybenzoyl)quinoline-2-carboxaldehyde) assay is based on binding of a nonfluorescent dye to primary amines of proteins in the presence of cyanide or thiols. The binding causes the dye to become fluorescent, and the fluorescence intensity is proportional to the amount of total protein. The methods described above vary in their ability to detect and quantify different proteins, are subject to interference from other molecules in the sample, and may require careful calibration; therefore, the particular choice of protein assay is highly dependent on the sample type.125 Nevertheless, these

4.2. Enzyme-Linked Immunosorbent Assay (ELISA)

The most common tool for protein detection is the immunoassay. The immunoassay was first introduced in 1960 to measure plasma insulin using radioactive labels.126 Due to safety concerns associated with the use of radioactive labels, including the need for special facilities and disposal of waste, alternative methods to replace radioactive labels were necessary. In 1971, an enzyme-based immunoassay was developed by two different groups independently in which the radioactive label was replaced by an enzyme label.127 Engvall and Perlmann developed the enzyme-linked immunosorbent assay (ELISA) for antibodies in rabbit serum using alkaline phosphatase,128 and van Weemen and Schuurs developed the enzyme immunoassay (EIA) for human chorionic gonadotropin in urine using horseradish peroxidase.129 In an ELISA, the enzyme label is used to amplify the signal by catalyzing the formation of many thousands of molecules of a detectable product. In this manner, each protein binding event is amplified. Currently, the enzyme-linked immunosorbent assay (ELISA) is the gold standard tool for protein detection and quantification. Commonly used enzymes include horseradish peroxidase (HRP), alkaline phosphatase (AP), and βgalactosidase. Many different substrates for these enzymes have been developed, and the products can be detected using various detection methods including colorimetric, fluorescent, chemiluminescent, and electrochemical detection. The main advantages of the ELISA include the ability to measure proteins quantitatively with relatively high sensitivity, in the pg/mL range, and with a wide dynamic range that spans 4 orders of magnitude. Although ELISAs are rather time consuming to carry out because they involve relatively long incubation times and extensive washing, they are easy to use. ELISAs are widely used for both clinical diagnostics and basic research. For example, ELISAs are commonly used in the clinic to detect C-reactive protein (CRP), a protein that plays an important role in activating the immune system.130 CRP levels in the blood are often elevated in subjects with inflammation; therefore, elevated CRP levels can be indicative of autoimmune disease or bacterial infection. Another example in which ELISAs are used in the clinic is for detecting antibodies against HIV in blood. HIV antibodies are detectable in the blood 3 months following infection. However, it has been shown that ultrasensitive immunoassays can detect HIV earlier than the 3month period. One example of such assay is the biobarcode assay, which is discussed in more detail in section 4.7.131 Approaches to improve protein detection are primarily based on improving the performance of ELISAs. 4.3. Enzyme-Linked Immunospot (ELISPOT)

The enzyme-linked immunospot (ELISPOT), which was developed in 1983, is used to detect secreted proteins by live cells in vitro.132 ELISPOT is conceptually similar to an ELISA. Briefly, antibodies are coated onto the surface of a microtiter plate. A sample containing cells that secrete proteins is added, and the target protein then binds to the antibodies. A second enzyme-labeled detection antibody is then added. The enzyme produces a colored precipitate. The assay is easy to perform on a large number of samples. This method is commonly used to detect cytokine secreting cells. ELISPOT assays have been H


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used to detect interferon gamma release for tuberculosis diagnosis.133 4.4. Western Blot

The Western blot, which was developed by three different groups independently between 1979 and 1981, is a widely used technique in life science research to identify specific proteins.134−136 In a Western blot, a sample is separated by size using gel electrophoresis, transferred to a solid membrane, blotted with a labeled antibody specific to the target protein, and then visualized.137 Western blots can be used for quantitative detection of proteins by using known standards. A major advantage of Western blots is the added layer of specificity, since proteins are initially separated by size. This additional specificity contrasts with ELISA, which only detects the label and does not provide any information on the molecular size of the protein. Even though the Western blot is rather time consuming and can only detect high protein concentrations, it is widely used due to its high specificity and simplicity, since it requires no complex instrumentation. A variation of the Western blot is the dot blot, which is similar to the Western blot but does not separate protein by electrophoresis. Recently, the single-cell Western blot was developed for measuring proteins with high sensitivity on a single-cell level. This method is described in section 6. The Western blot is most widely used for basic research and is commonly used to detect protein isoforms, truncated proteins, post-translational modifications, and antibodies. Western blots are also used in the clinic. For example, HIV diagnosis is typically a two-step process in which the first step is detecting antibodies against HIV in the blood using ELISA and the second step is confirming the ELISA result using a Western blot.138,139 This combined approach leads to 99% clinical sensitivity and specificity.

Figure 7. Protein microarrays. (a) In a planar microarray, capture antibodies specific to different proteins are immobilized at known and predefined positions for multiplexed detection. Target protein binds to the capture antibody and then to a labeled detection antibody. Signal intensity is correlated with the presence and amount of the target protein. (b) In a reverse phase protein microarray, the sample is deposited onto the surface of the microarray and the target protein is detected by probing with an antibody. Signal intensity is correlated with the presence of the target protein. Modified and reprinted with permission from ref 117. Copyright 2015 Nature Publishing Group.

ProtoArray from ThermoFisher. Protein microarrays can be used to study protein−protein interactions and enzymatic activity and for biomarker discovery.141 One of the earliest implementations of protein microarrays was for analyzing the biochemical activity of yeast proteins, including identifying proteins that interact with calmodulin and phospholipids and analyzing protein kinases.142,143 More recently, protein microarrays were used for high-throughput detection of antibodies against Plasmodium falciparum (Pf) in plasma before and after malaria season.144 Additionally, protein microarrays have been used for analyzing protein interactions with other molecules including both nucleic acids145 and small molecules.146 Protein microarrays are also used for clinical diagnostics.147 One example is the Meso Scale Discovery (MSD) system. Another example is the SOMAscan assay, which uses a nucleic acid microarray for protein detection. These methods are further described in section 6.

4.5. Protein Microarrays

Protein microarrays are used for multiplexed detection of proteins using reagents immobilized on a planar surface typically made of silicon, glass, or nitrocellulose. Reagents can be immobilized onto the surface of the microarray by various approaches including contact printing or in situ synthesis of reagents.140 There are two types of protein microarrays. The first is based on a sandwich immunoassay (Figure 7A) in which capture antibodies are immobilized onto the surface of the microarray. The target protein first binds to the capture antibodies, labeled with a detection antibody, and detected. This type of microarray can be used for multiplexed detection of proteins by immobilizing antibodies specific to different proteins at known and predefined positions. In the second type of microarray, also known as a reverse phase array (Figure 7B), a sample is deposited onto the surface of the microarray and the target protein is detected by probing with an antibody. Reverse phase arrays are commonly used to detect protein phosphorylation levels or antibodies. In protein microarrays, the detected signal is usually fluorescent, chemiluminescent, electrochemiluminescent, or colorimetric. Typically, the binding kinetics of protein microarrays are less favorable than the binding kinetics of solution-based assays using microspheres since the proteins must diffuse to the surface of the microarray in order to bind to the capture reagent. This mass transport limitation may result in reduced sensitives for protein microarrays. Protein microarrays are commonly used for protein detection and are available commercially. One example is the

4.6. Flow Cytometry

Flow cytometry was first introduced in the 1960s and is used to separate cells based on size and protein surface markers. In flow cytometry, fluorophore-labeled antibodies bind to specific cell-surface markers. The cells are then flowed through a flow channel. Lasers are used to excite the fluorophores, and the emission is measured at two different angles, providing the ability to differentiate one cell at a time based on size and surface markers. The main advantage of flow cytometers is their high throughput, namely, their ability to analyze thousands of cells per second. Another advantage is the ability to sort using fluorescence-activated cell sorting (FACS) for further downstream analysis of cells. Flow cytometry is commonly used by immunologists, particularly to detect cellsurface proteins known as clusters of differentiation (CDs). For example, analysis of cell-surface markers using flow cytometry was instrumental for characterizing B-cell lineage development. Hardy et al. showed that progenitor and precursor B cells in the bone marrow are a complex mixture of cells at different stages of development.148 Flow cytometry also has applications in clinical diagnostics of immunodeficiency and hematopoietic disorders based on cell-surface protein markers.149 In 2017, the FDA approved for the first time a flow-cytometry-based assay for blood cancer diagI


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nostics.150 This assay detects several cell-surface markers in peripheral blood, bone marrow, and lymph node samples for diagnosis of leukemia, non-Hodgkin lymphoma, multiple myeloma, myelodysplastic syndrome (MDS), and myeloproliferative neoplasms (MPN). Other than cells, flow cytometers can also be used to detect specific proteins using microspheres in a sandwich assay format. Microspheres are coated with specific antibodies, incubated with the target protein, and then labeled with a second fluorescently labeled detection antibody. Flow cytometry is becoming increasingly popular for protein quantification due to the high multiplexing capabilities. Fluorescent labels are now being replaced with metal ions in a new technique known as mass cytometry. These new techniques are discussed in more detail in section 6. The main limitation of flow cytometry is high cost, the need for a skilled operator, and relatively complex instrumentation.

cleotides complementary to the oligonucleotides connecting the two antibodies are added and two restriction sites are generated for digestion. The resulting DNA strands are circularized by a second ligation and then amplified by RCA. The amplified products are then detected by hybridization with fluorescently labeled probes and counted using a microfluidic device. An alternative to PLA is the proximity extension assay (PEA), in which the DNA ligase is replaced by a DNA polymerase. PEA aims to reduce the interfering effects of DNA ligase in biological samples.158 Multiplexed PLAs have been developed by using probes with different sequences and lengths, allowing for development of a 24 plex assay.159 Both PLA and PEA provide sensitive methods for protein detection with subfemtomolar sensitivity using a low sample volume. These assays can be used in solution-based assays to measure protein levels and detect protein−protein interactions. They can also be used for protein analysis in tissue samples. Nucleic acid assays have been used for highly sensitive protein assays with detection limits in the attomolar range.160,161 Recently, new methods have been developed for DNA-based protein detection, including use of DNA nanoswitches,162 digital droplet PCR,163,164 and incorporation of nanoparticles.165 Another method for ultrasensitive protein detection is the biobarcode assay developed in the Mirkin laboratory (Figure 9).160,161,166,167 First, antibody-functionalized magnetic particles bind to the target protein. Then gold nanoparticles, which are functionalized with both detection antibodies and oligonucleotide barcodes, bind to the target protein to form a sandwich complex. The oligonucleotide barcodes are then released. Prior to detection, the biobarcodes can also be amplified by PCR. The biobarcodes hybridize to probes on a microarray that are complementary to one-half of the biobarcode sequence. Following hybridization to complementary probes on the microarray, nanoparticles functionalized with oligonucleotides complementary to the other half of the biobarcode sequence hybridize. Silver amplification is used to enhance the signal, and gray spots can be detected using a Verigene instrument. The assay results in high sensitivity in the attomolar range. The biobarcode assay has been used for the sensitive detection of prostate specific antigen (PSA) in the blood of patients who have undergone radical prostatectomy.168 Following radical prostatectomy, PSA is usually not detectable using a conventional ELISA. Since increasing PSA levels can be indicative of recurrence, high-sensitivity assays such as the biobarcode assay can be used to detect recurrence earlier. Multiplexed detection can be achieved by using gold nanoparticles each functionalized with a unique biobarcode sequence (Figure 9).111 These unique biobarcodes hybridize to complementary probes at predetermined positions on a microarray. Another method that uses a DNA microarray for multiplexed protein detection is the SOMAscan assay, which is further discussed in section 6.2.1.

4.7. Proximity Ligation Assay (PLA) and other Nucleic Acid-Based Detection Methods

An approach for signal generation and amplification is to convert protein binding into a nucleic acid signal.151,152 The main advantage is that highly sensitive detection of nucleic acids is possible because DNA can be amplified. By combining the benefits provided by antibodies and the amplification advantage provided by PCR, immuno-PCR was introduced and enabled protein detection with greater sensitivity than the conventional immunoassay. In this approach, the target protein binds to capture antibodies immobilized on a surface. A DNA−antibody conjugate then binds to the target protein. The DNA is amplified by PCR and then detected by gel electrophoresis.153 Further improvements to immuno-PCR have been realized by using a sandwich assay consisting of a capture antibody and a detection antibody that is conjugated to an oligonucleotide label, amplifying the oligonucleotide label and quantifying the label using qPCR.154 Another method called the proximity ligation assay (PLA) is used for protein detection (Figure 8).155,156 In PLA, two biotinylated

Figure 8. Proximity ligation assay (PLA). In PLA, the sample is first incubated with antibodies conjugated to oligonucleotide probes (1). Components for ligation and detection are added to the sample (2), and quantitative PCR is used for detection (3). Modified and reprinted with permission from ref 156. Copyright 2004 National Academy of Science.

antibodies bind to different epitopes of the target protein analyte. Two different streptavidin-labeled oligonucleotides are bound to each of the two biotinylated antibodies. PLA is designed to give a signal only when a protein is present due to the proximity of the epitopes within the target protein. When both oligo-labeled antibodies bind to the protein, a connector oligonucleotide can hybridize to both of the oligonucleotides, a ligation step is performed, resulting in a DNA template that can be amplified and quantified using qPCR.156 A variation of PLA utilizes rolling circle amplification (RCA) for singlemolecule detection using digital PCR that enhances the sensitivity of the method.157 More specifically, two oligonu-

4.8. Mass Spectrometry

Mass spectrometry is a common tool for proteomics analysis. It is probably the most comprehensive method for analyzing complex mixtures because it provides a full profile of the protein composition. Conventional tandem mass spectrometry digests proteins into peptides, and these peptides can then be used to derive the sequence of the original protein molecule. This strategy, known as “bottom-up” proteomics, provides invaluable information about proteins present in complex J


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Figure 9. Biobarcode assay. Target protein binds to magnetic microparticles (MMPs) functionalized with capture antibodies. Each population of nanoparticles (NP) is functionalized with unique biobarcodes and detection antibodies specific to a target protein. Nanoparticles bind to the target protein. Biobarcodes are then released and hybridize to probes complementary to one-half of the sequence. Gold nanoparticles then hybridized to the other half of the sequence. Silver is used to enhance the signal, and the signal intensity is measured. Modified and reprinted with permission from ref 166. Copyright 2006 Nature Publishing Group.

samples.169 However, since the same peptide sequence may be present in many different proteins, it is often challenging to identify different proteins, such as those that arise from different gene product and splice variants. Another strategy that may overcome this challenge is “top-down” proteomics, in which whole, intact proteins are identified directly using tandem mass spectrometry. This approach enables analysis of different types of proteins; however, it is still challenging to analyze low-abundance proteins, and quantification is also difficult. Additional information on mass spectrometry proteomics techniques and advances is provided in other reviews.12,13,170−173 4.9. Lateral Flow Assay (LFA)

Lateral flow assays (LFAs) are commonly used for clinical diagnostics and point of care detection of proteins in various sample matrices including blood and urine.174,175 The most commonly used assay format is the sandwich assay. An LFA assay consists of several zones, and a typical configuration is shown in Figure 10. First, the sample is added to the sample application pad. The sample then moves toward the conjugate pad, which houses an antibody conjugated to a label. Commonly used labels for LFAs are latex particles or gold nanoparticles, which can form colored precipitates. Next, the sample, which contains the protein bound to a labeled antibody, moves to a band that contains immobilized capture antibodies. A sandwich complex is then formed. Often there is a second control band, which is immobilized with antibodies that capture the labeled antibody, to ensure the test is performing correctly. Excess reagents then migrate past these bands and are trapped in the wick or absorbent pad. The signal intensity of the band is proportional to the concentration of the target protein. The results can be visualized with the naked eye or with a reader. Usually LFAs provide qualitative or semiquantitative protein measurements. One limitation is that when very high levels of protein target are present in the sample, the antibody binding sites become saturated. This results in a high-dose hook effect, which produces a lower signal intensity than expected. LFAs are important for clinical diagnostics. A major advantage of LFAs is that they are cheap and rapid, with results provided in minutes, do not require complex instrumentation, and are compatible with point of care applications. The most common use is the pregnancy test,

Figure 10. Sandwich lateral flow assay (LFA). (Top) In a sandwich LFA, the sample is first added to the sample application pad and then migrates toward the conjugate pad, which houses antibodies conjugated to a label. Protein binds to the labeled antibodies, and sample then moves to a band that contains immobilized capture antibodies. Sandwich complex is then formed. Control band is immobilized with antibodies that capture the labeled antibody to ensure the test is performing correctly. Excess reagents then migrate past the bands and are trapped in the absorbent pad. (Bottom) Signal intensity of the band is proportional to the concentration of the target protein. Reprinted with permission from refs 175 and 177. Copyright 2016 Elsevier and 2009 MDPI.

which measures the presence of the protein human chorionic gonadotropin (hCG) in urine.176 4.10. Surface Plasmon Resonance (SPR)

Surface plasmon resonance (SPR) is a label-free method that can be used for protein detection. SPR occurs when conduction electrons undergo oscillations at metallic surfaces upon exposure to light.178,179 SPR detection of proteins is based on immobilization of an affinity reagent onto a metal surface. The affinity reagent then specifically binds to the protein of interest in the sample. The binding event causes a measurable change in the refractive index. SPR intensity depends on the dielectric properties of the medium as well as K


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size, composition, and shape of the metallic structure.180 Additionally, SPR is highly dependent on the angle of incidence of light, the wavelength of light, and the refractive index of the sample. SPR is commonly used to study the binding kinetics of protein−protein interactions, such as binding of proteins to antibodies.181 Additionally, SPR has been used to detect live malignant cells.182 Gold nanoparticles conjugated to antiepidermal growth factor receptor (antiEGFR) antibodies were added to cultures of nonmalignant and malignant cells. The anti-EGFR antibody-conjugated nanoparticles bound to the surface of the malignant cells with substantially higher affinity than to the nonmalignant cells and were detected by SPR. More recently, SPR has been used to detect exosomes, nanometer-sized extracellular vesicles, based on surface protein markers.183 SPR is advantageous due to its ability to generate quantitative measurements with a lack of labeling variations associated with optical labels such as fluorescent dyes. Localized surface plasmon resonance (LSPR) of metal nanostructures can be used to enhance the signal.184,185 Additionally, nanoparticle labels on detection antibodies in a sandwich assay format can also be used to enhance the signal.186,187

slide, enable high-throughput analysis of many samples. Tissue microarrays have been used for the Human Protein Atlas for analysis of protein expression in normal and cancer human tissue.1,192,193 Live cell imaging is becoming more popular to study protein localization and dynamics in live cells. Since antibodies cannot penetrate the cell membrane, proteins are genetically tagged with fluorescent molecules, such as green fluorescent protein (GFP). Other systems that are based on SNAP, CLIP, and Halo tags have also been used.194 The study and detection of proteins in the context of their cellular environment can be enhanced by improvements in fluorescence microscopy. Super-resolution microscopy can resolve features less than the diffraction limit of light, which is about 200 nm in the lateral direction and 500 nm in the axial direction. These techniques include stimulated emission depletion (STED), photoactivation localization microscopy (PALM), stochastic optical reconstruction microscopy (STORM), structured illumination microscopy (SIM), saturated structured illumination microscopy (SSIM), ground state depletion (GSD), and reversible saturated optical (fluorescence) transitions (RESOLFT) and have been thoroughly reviewed.195−198

4.11. Optical Imaging

5. CHALLENGES WITH PROTEIN DETECTION The performance of a protein detection technique is assessed by various parameters including sensitivity, specificity, dynamic range, multiplexing capabilities, reproducibility, ease of use, and cost. Currently, a major analytical challenge with protein detection techniques is measuring multiple proteins, which are present in complex biological samples, with high sensitivity and a wide dynamic range. In this section, we will discuss some of the current challenges, including sensitivity, specificity, and multiplexing capabilities. High-sensitivity measurements of proteins are particularly important for many applications. One example is the study of single cells. Single-cell transcriptomics has been widely used to study heterogeneity between different cells and for understanding important biological processes such as stem cell differentiation and cancer. However, our understanding of protein levels and dynamics in single cells is limited. Additionally, the relationship between mRNA levels and protein levels on a single-cell level has been studied164,199−201 but is still not fully understood. Another example is measurement of blood protein biomarkers for clinical diagnostics, including disease diagnosis, monitoring, and treatment. It is particularly challenging to detect lowabundance proteins in the presence of high-abundance proteins in the blood. Less than 10% of plasma proteins have been quantitatively measured due to limitations in analytical sensitivity.202,203 Therefore, high-sensitivity measurements of proteins are critical. The sensitivity, or detection limit, is typically calculated based on the background signal of the assay.204,205 The background signal is due to nonspecific binding of the affinity reagents, labeling reagents, and components in the biological sample with each other or with the surface. The detection limit is usually calculated based on measuring a pure protein in a buffer solution. However, many biological samples contain interfering substances that may increase the background signal, and therefore, it is important to assess the detection limit in the biological sample matrix. To reduce nonspecific binding arising from sample components, sample separation techniques can be

The subcellular localization of proteins is important for function due to the unique chemical environment and proximity to other interacting molecules. Delocalization of proteins may result in a lack of function and disease. Therefore, the spatial distribution of proteins in their native cellular context is critical for understanding their function. Optical imaging-based techniques are instrumental for studying proteins in their native cellular environment. Immunohistochemistry (IHC), first developed in 1941,188 is used to detect proteins and their cellular localizations in tissues and cells. Traditionally, a sample is first fixed or frozen; then the antibody binds to the target protein and is then labeled to produce a colored product that can be optically detected. Antibody binding can be observed either by using a fluorophore or an enzymatic reaction that produces a colored product. IHC is a widely used technique for protein detection, particularly in clinical diagnostics, where it is used for disease diagnosis, prognosis, and treatment.189 Another similar technique, immunocytochemistry (ICC), is used to analyze proteins in cultured cells or cells in which the extracellular matrix has been removed. Since sample preparation is different for IHC and ICC, a given antibody may not be compatible with both methods since the protein epitope may not be fully exposed and cannot bind to the antibody. For IHC, a process known as antigen retrieval is commonly used to unmask the epitope by heating or digesting the sample with proteases. In many cases a combination of both heating and digestion is used. This process unmasks the protein epitope and allows the antibody to bind to the protein.190 The main advantage of these techniques over other protein detection methods is the ability to correlate the presence of a protein with its location in a tissue or cell. This feature is very important for studying cell function in normal and pathological tissues and for differentiating between healthy and diseased tissue. Recently, Thul et al. used immunofluorescence microscopy to map the subcellular location of approximately 12 000 human proteins.191 Tissue microarrays, in which hundreds of different tissue samples are assembled on a single L


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employed including desalting, filtration, and affinity depletion of other interfering molecules. As a result, the particular type of sample is also important since different samples may contain different compositions of interfering substances. Another way to overcome nonspecific binding is by optimizing buffer components, such as adding blockers. An important parameter in protein detection is conversion of the target protein binding with the affinity reagent into a measurable signal. Detection of a specific protein in a complex biological matrix is often challenging. Biological samples contain many protein and nonprotein analytes that may interfere with detection of the target protein. Some nonprotein target analytes may be present at considerably higher levels than the target protein, further complicating detection. Additionally, nontarget proteins with similar structures may bind to the affinity reagents and interfere with accurate quantification. High-affinity reagents that are specific to the target protein and do not bind to other nonanalyte molecules are necessary to enhance specificity. Another challenge arises for some types of assays in which an affinity reagent may specifically bind to a target protein in a certain method but not in another. For example, proteins are detected in their native form for sandwich immunoassays and flow cytometry, but they are denatured in Western blots and immunohistochemistry. Due to differences in protein conformation and epitope structure, affinity reagents that target the same protein may not be usable across different methods. Therefore, detecting a specific protein target in a biological sample is often quite challenging. Another challenge with protein detection methods is multiplexing. It is advantageous to simultaneously detect multiple proteins in a single sample for increased throughput, lower cost, reduced sample volume, and faster assay time. A major challenge with multiplexed assays is cross-reactivity between components in the assay, including affinity reagents and other proteins, which often complicates detection and accurate quantification. In many samples, the concentrations of different proteins in a biological sample can span several orders of magnitude. Therefore, it is advantageous to have a wide dynamic range to measure both high- and low-abundance proteins simultaneously in a single sample. A wide dynamic range is also useful for measuring a single protein in different biological samples since some samples may have substantially different levels of a given protein. Multiplexed assays also often reduce the sample volume requirements. This feature is particularly important when sample volume is limited, for example, when measuring proteins in cerebrospinal fluid, tumor tissue samples, or blood obtained from infants. Therefore, a major analytical challenge is measuring multiple proteins in a complex biological sample with both high sensitivity and a wide dynamic range. Finally, other analytical parameters of the assay, including precision, accuracy, and reproducibility, are important. A final consideration is whether a simple binary result, such as the presence or absence of the target protein, is sufficient or if quantitation is required. For detailed discussion of analytical parameters of immunoassays, the reader is referred to a literature review.206

measuring multiple proteins simultaneously with high sensitivity. In this section, we will review techniques that are currently being implemented to overcome limitations in both sensitivity and multiplexing. 6.1. Ultrasensitive Methods for Protein Detection

6.1.1. Meso Scale Discovery (MSD). The Meso Scale Discovery System (MSD) is based on an electrochemiluminescent (ECL) sandwich immunoassay that can measure up to 10 different proteins simultaneously (Figure 11).207

Figure 11. Meso scale discovery (MSD) assay. (a) Generation of electrochemiluminescence (ECL) based on oxidation of ruthenium at an electrode in the presence of tripropylamine (TPA). (b) ECL ruthenium label used to modify antibodies. (c) MSD assay plate consisting of an injection-molded plate top and a bottom plate consisting of screen-printed carbon ink electrodes. (d) Well containing electrodes and antibodies. Each well contains 10 spatially defined positions that house capture antibodies specific to different proteins for multiplexed detection. (e) Sandwich immunoassay with ECL detection. (f) Sector PR 400 plate reader. Reprinted with permission from ref 207. Copyright 2007 Elsevier.

Electrochemiluminescence (ECL) involves a chemical reaction that generates intermediates, which undergo an electron transfer reaction at the electrode surface to produce excited states that emit light.208−211 Inorganic molecules, such as tris(2,2′-bipyridine)ruthenium(II) (Ru(bpy)32+), are widely used to generate ECL. Due to its low molecular mass, many Ru(bpy)32+ molecules can be attached to an antibody without hindering the antibody’s ability to bind to the target protein. Additionally, when a photon is emitted, the ground state is regenerated, and therefore, a single label can undergo many reaction cycles to produce numerous photons. Overall, signal transduction using ECL is advantageous for several reasons including high sensitivity, wide dynamic range, simplicity, and stability. A biological sample may contain molecules that are autofluorescent and increase the background signal; therefore, a major advantage of ECL over fluorescence detection is the lack of a need for an excitation source. This results in lower

6. EMERGING METHODS FOR ULTRASENSITIVE AND HIGHLY MULTIPLEXED MEASUREMENTS Emerging new techniques have been developed to overcome the challenges discussed in the previous section. A major analytical challenge with protein detection techniques is M


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Figure 12. Single-molecule array (Simoa) immunoassay. (a) Single protein molecule is first captured on beads, which are functionalized with capture antibodies. Protein is then labeled with a biotinylated detection antibody and streptavidin-ß-galactosidase. (b) Excess beads are added to a sample containing a low concentration of protein so that only one or zero protein molecules bind to each bead. In the presence of enzyme substrate, beads are then loaded onto an array of femtoliter-sized wells such that only one bead can fit per well. Array is sealed with oil, and fluorescence images of the array are then obtained. (c) Scanning electron micrograph image of a microwell array containing beads inside wells of femtoliter volume. (d) Fluorescence images of a microwell array following generation of signal from a single enzyme molecule. Reprinted with permission from ref 219. Copyright 2010 Nature Publishing Group.

background signals by eliminating detection of fluorescence that is due to other components in the sample. Enhanced ECL signals can also be generated, and several reviews have been written on the topic.212−216 In the MSD assay, which is based on ECL detection, capture antibodies are first screen-printed onto the bottom of a 96- or 384-well plate at predetermined positions. After binding of the target protein to the capture antibody, a second detection antibody that is labeled with ruthenium binds to the target protein. When the ruthenium complex is placed at an oxidizing electrode in the presence of tripropylamine (TPA), an ECL signal is generated. The MSD assays have sensitivities in the low pg/mL range and a wide dynamic range of 4 orders of magnitude.217 MSD assays are commonly used to measure protein biomarkers in blood for clinical applications. 6.1.2. Single-Molecule Arrays (Simoa). Digital ELISA using single-molecule arrays (Simoa), which is being commercialized by Quanterix Corp., is an ultrasensitive method for protein detection using a bead-based sandwich ELISA.218 In a conventional sandwich ELISA, the fluorescent product of the enzyme−substrate reaction diffuses into a large volume of approximately 50−100 μL. Therefore, millions of enzyme-labeled immunocomplexes are required to generate a measurable fluorescent signal above the background. In digital ELISA, the fluorescent product of the enzyme−substrate reaction is confined into femtoliter-sized wells, which are termed single-molecule arrays (Simoa).219 Using this approach, the presence of a single-enzyme molecule can be detected.220 In a Simoa immunoassay (Figure 12), antibody-coated capture beads are added in excess to a sample containing low concentrations of target protein molecules. On the basis of Poisson statistics, either one or zero target protein molecules will bind to each bead. To illustrate this concept, a 100 μL amount of blood sample with 1 fM of the target protein contains approximately 60 000 protein molecules. If 500 000

antibody-coated beads are incubated with the blood sample, most of the beads will bind zero protein molecules while a small number of beads will bind one protein molecule. A negligible number of beads will bind more than one protein molecule based on the Poisson distribution. The beads are then incubated with a biotinylated detection antibody and streptavidin-ß-galactosidase, forming an enzyme-labeled immunocomplex. The beads are then loaded onto an array of femtoliter-sized wells, in which each well is physically able to hold exactly one bead. The wells are filled with a fluorogenic substrate and sealed with oil. Beads containing a single immunocomplex will result in enzymatic conversion of many substrate molecules into product. Because the fluorescent product is confined to an extremely small volume of approximately 50 fL, a high local concentration of fluorescent product results that is easily detected with an imaging detector. Hundreds of thousands of wells are interrogated simultaneously, and the ratio of beads containing an enzyme label to the number of total beads in wells corresponds to the target protein concentration in the original sample. In this manner, single-protein molecules can be detected using a digital readout. A wider dynamic range beyond the digital range can be obtained by using the average fluorescence intensity of the active beads. This results in a dynamic range that spans over 4 orders of magnitude.221 Simoa immunoassays have been developed for multiplexed detection of six different proteins using dye-encoded beads.222 Simoa immunoassays have been used for ultrasensitive detection of biomarkers with applications in neurology,223 cardiology,224 oncology,225,226 and immunology.227 6.1.3. Single-Molecule Counting (SMC). The singlemolecule counting (SMC) platform, also known as the Erenna immunoassay system from Singulex Inc., is capable of highsensitivity protein measurements with detection limits in the sub pg/mL range.228,229 SMC is based on a traditional sandwich immunoassay, in which a capture antibody N


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allowed to settle into a well. The cells are then lysed. The proteins are loaded electrophoretically into the surrounding gel, and all of the Western blot steps including electrophoresis, blotting, and probing are performed. This technique has several advantages including the ability to assay low volumes and low concentrations of proteins. An additional advantage is the ability to integrate antibody probing with gel electrophoresis. This method allows detection of multiple protein isoforms as well as truncated proteins, which may be of importance for disease detection. Using this approach, multiplexed detection of 12 different proteins has been accomplished.232

immobilized on the surface of either a microtiter plate or magnetic beads binds to the target protein. A second detection antibody that is conjugated to a fluorophore then binds to a different epitope on the target protein, forming a sandwich complex. Following several washing steps, the fluorescently labeled detection antibody is eluted from the immunocomplex into a small volume. The eluate is detected using a laser in a small interrogation area. As the fluorescently labeled detection antibodies flow through the interrogation area, they are detected as positive or negative events using a confocal imaging system. The emitted photons are counted over defined time intervals, and if the photon count is sufficiently higher than the background, the signal is considered positive. By counting the number of positive events, a digital readout is obtained. At higher protein concentrations, an analog mode is used in which the total number of photons is used. This provides a wide dynamic range of approximately 4 orders of magnitude. The original system was designed for single-plex assays; however, recently, a three-plex assay was developed using three labels and three different lasers.230 6.1.4. Single-Cell Western Blot. A sensitive protein detection method for one specific application is the single-cell Western blot (Figure 13).231,232 Many protein detection

6.2. Highly Multiplexed Methods for Protein Detection

6.2.1. SOMAscan Assay. Oligonucleotides containing modified nucleic acid side chains can bind to proteins with high affinity. Various side-chain modifications for deoxyuridine triphosphate (dUTP) have been developed to enhance binding affinity. These modifications mimic the hydrophobic amino acid residues tryptophan and phenylalanine, which are found in many protein−protein binding regions. The modified aptamers can achieve high affinities, with dissociation constants of less than 1 nM. These high-affinity reagents are termed slow offrate-modified aptamers, abbreviated as SOMAmers.234,235 On the basis of these SOMAmers, SomaLogic developed the SOMAscan assay for highly multiplexed protein detection. The principles of the assay are shown in Figure 14. For protein detection, SOMAmers are designed with three tags: a fluorophore, a photocleavable linker, and biotin. The SOMAmer reagents first bind to beads functionalized with streptavidin via biotin−streptavidin interaction. Then the beads are incubated with the sample that contains the target protein, and the target protein binds to the SOMAmer reagent attached to the beads. The unbound proteins are washed away, and then the proteins bound to the SOMAmer are tagged with biotin. Ultraviolet light is then used to break the photocleavable linker and release the SOMAmer reagent. Some of the released SOMAmer reagents are associated with a biotinylated target protein. A polyanionic competitor is then added to further reduce nonspecific binding. Streptavidinfunctionalized beads are then used to recapture the SOMAmer-biotinylated target protein complex. Additional washing steps remove nonspecifically bound SOMAmers. The SOMAmer reagent is then released in a denaturing buffer and hybridized to an array of single-stranded complementary DNA probes. Fluorescence intensity is then related to the protein levels in the sample. The SOMAscan assay has been used for biomarker discovery, primarily in biological samples such as blood.236 The main advantage of the SOMAscan assay is its high multiplexing capabilities. SomaLogic has generated SOMAmers specific to approximately 1300 human proteins with sensitivities in the low-picomolar to high-femtomolar range.237,238 A major goal of the company is to develop additional SOMAmers that recognize many different proteins to enable proteomics-level measurements. However, since the technology is still in its early stages, additional analytical variables must be determined, such as cross-reactivity and assay variability.239 Nevertheless, this platform enables highly multiplexed protein measurements with sufficient sensitivity for unbiased protein measurements. 6.2.2. Luminex. The Luminex xMAP system is based on a sandwich immunoassay format that incorporates flow

Figure 13. Single-cell Western blot. First, single cells settle into wells in a microwell array made of a photoactive gel. Dimensions of the wells are such that only one cell can fit inside each well. Cells are then lysed with a denaturing RIPA buffer. After polyacrylamide gel electrophoresis (PAGE), the proteins are immobilized onto the gel with UV light followed by probing with a primary antibody (1° Ab) and then secondary labeled antibody (2° Ab*). Fluorescent images of the array are obtained to detect the target protein. Chemical stripping and reprobing enables detection of multiple proteins. Modified and reprinted with permission from ref 233. Copyright 2014 Nature Publishing Group.

methods use antibodies to capture a specific protein. While antibodies are enabling for protein detection, limitations exist due to antibody specificity and cross-reactivity with other similar proteins. The Western blot, one of the most commonly used techniques for protein detection, first separates proteins based on molecular mass using gel electrophoresis prior to antibody probing and is thus more specific for the target protein. In the traditional Western blot, thousands of cells are needed to overcome the background signal. There is increasing interest in analyzing proteins on the single-cell level, for example, for identification of rare circulating tumor cells (CTCs) in the blood. Microfluidic devices offer an advantage by reducing the amount of required sample, facilitating separation of rare CTCs from the more abundant red and white blood cells. However, a major challenge is downstream analysis of the isolated CTCs on a single-cell level. The single-cell Western blot addresses some of these limitations. The device is comprised of a microscope slide and a layer of polyacrylamide gel that is patterned with an array of microwells, in which each well can physically hold only a single cell. A single cell is loaded onto the microwell array and O


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Figure 14. SOMAscan assay. (a) SOMAmers, which are functionalized with a fluorophore, photocleavable linker, and biotin, are immobilized onto streptavidin (SA)-coated microspheres and incubated with a sample. (b) Target protein binds to the SOMAmer reagents. (c) Target protein is tagged with biotin. (d) UV light is used to cleave the photocleavable linker on the SOMAmer and release the SOMAmer−protein complex from the microspheres. (e) Addition of a buffer containing a polyanionic competitor disrupts nonspecific binding. (f) SOMAmer−protein complexes are captured on a new set of streptavidin-coated microspheres via the biotin on the target protein. (g) Denaturing buffer is used to release the SOMAmers from the microspheres. (h) SOMAmers hybridize to complementary probes on a microarray and are detected by fluorescence. Fluorescence intensity is correlated to the target protein concentration. Reprinted with permission from ref 236. Copyright 2014 Elsevier.

Figure 15. Mass cytometry (CyTOF). Antibodies labeled with heavy isotopes bind to the target proteins. Each cell is nebulized and introduced into an inductively coupled plasma (ICP) and ionized. Abundant ions are removed, and remaining heavy elements are analyzed. Signal of each isotope label is correlated to the presence of the target protein. Reprinted with permission from ref 244. Copyright 2012 Cell Press.

cytometry as the detection platform. Luminex assays can achieve high multiplexing capabilities of 100 analytes by entrapping different ratios of dyes into microspheres to obtain optically distinct populations. Each optically distinct population of microspheres is attached to capture antibodies specific to a given protein. Following binding of the target protein, a second antibody conjugated to phycoerythrin (PE) binds to the protein. The microspheres are then analyzed using a flow cytometer to detect the internal fluorescent labels to identify the microsphere’s specificity and measure the PE emission

intensity corresponding to the amount of target protein bound to the bead. A major advantage of the Luminex system is its high-multiplexing capabilities. However, commercially available kits typically do not exceed simultaneous measurements of more than 30 proteins due to cross-reactivity. The sensitivity of Luminex assays is generally between 1 and 10 pg/mL range, and the dynamic range spans approximately 3−4 orders of magnitude. The Luminex system is widely used to measure proteins for various research and approved in vitro diagnostics (IVDs) applications including infectious diseases, human P


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leukocyte antigen (HLA) detection, and more.240 Another similar platform is the cytometric bead array (CBA), a beadbased sandwich immunoassay based on flow cytometry, that is used for multiplexed detection of proteins.241,242 6.2.3. Mass Cytometry (CyTOF). Mass cytometry was developed in 2009 to allow for protein detection on a singlecell level.243 Mass cytometry, also known as CyTOF, combines the principles of flow cytometry and atomic mass spectrometry, enabling analysis of 40 parameters per cell with very high throughput.244,245 The work flow of single-cell mass cytometry (Figure 15) includes coupling antibodies to a stable nonradioactive rare-earth metal isotope. The labeled antibodies bind to the target protein of interest. The labeled cell is then nebulized into a droplet and introduced into an inductively coupled argon plasma (ICP). Once the cell enters the plasma, the sample is vaporized and the cellular material in the droplet is ionized, including the abundant isotopes in the cell such as carbon, nitrogen, and oxygen. The ionized sample is passed through a mass filter, which allows only the heavy reporter ions through, and these ions are analyzed by the mass spectrometer. The mass spectrometer then measures the composition of the metal isotopes. This number is correlated to the abundance of antibody bound targets in the original sample. Mass cytometry has been used for various applications. Examples include single-cell analysis of immune and drug responses246 and analysis of T cells in healthy and HIV-infected subjects.247 Recently, metal isotopes have been used for multiplexed detection of proteins in tissue samples.248,249 The main advantage of mass cytometry is the ability to overcome the need for fluorophores by labeling antibodies with rare-earth heavy metals. The use of heavy metals provides an advantage for multiplexed detection since there are about 100 isotopes that are potentially available for mass cytometry and only 18 fluorophores for flow cytometry due to the spectral overlap of fluorophores. Additionally, the signal from the heavy rare-earth labels has low background because there are no rare earths present in biological samples. Therefore, the measured signal is highly specific to the rare-earth labels. This contrasts with optical detection of fluorophores, in which the biological sample may contain molecules that are autofluorescent and interfere with detection of the fluorescent label. Another advantage of mass cytometry is its very high throughput, with an ability to assay millions of single cells. The main disadvantages of the method are that it can provide only relative quantification and has low sensitivity. Several other disadvantages are a lack of ability to sort the cells since they are vaporized, challenges with quality control, statistical analysis of complex data, and limited availability of commercial rare-earth metal-labeled antibodies.

7.1.1. Upconverting Nanoparticles. Lanthanides exhibit anti-Stokes luminescence or upconversion luminescence.250 Upconversion is a nonlinear optical process in which absorption of two or more photons leads to emission of a photon at a shorter wavelength. Upconverting nanoparticles can be used in a sandwich assay for protein detection, in which the detection antibody is labeled with an upconverting nanoparticle. The label is excited in the infrared region and emits in the visible region for detection (Figure 16).251

Figure 16. Upconverting nanoparticles. Absorption of near IR light (NIR) and energy transfer lead to upconverted blue, green, or red emissions. Reprinted with permission from ref 251. Copyright 2009 John Wiley and Sons.

Upconverting nanoparticles have several favorable optical properties including high stability and quantum yield. Additionally, background arising from absorbing molecules in a biological sample is low since most biomolecules in a sample absorb in the UV region and not in the IR region. Upconverting nanoparticles have been used as optical labels for multiplexed detection and optical encoding.252−254 Several reviews have been written on their properties and applications.255−262 7.1.2. Photoelectrochemical Detection (PEC). In photoelectrochemical (PEC) signal transduction, light is used to excite photoactive species, which produce a measurable electrical signal (Figure 17). Since the excitation source (optical) is different than the detected signal (electrical), PEC-

7. ADVANCES FROM THE LITERATURE 7.1. Advances in Labeling and Signal Detection

A major consideration for protein assays is converting the binding of the target protein to an affinity reagent into a measurable optical, electrochemical, or mechanical signal. In general, enhancing the signal leads to higher sensitivities. Various approaches to enhance the signal of labeling reagents have been developed. In this section, we discuss several novel and promising approaches for labeling and signal detection, including upconverting nanoparticles, photoelectrochemical detection, optical ring resonators, and surface-enhanced Raman scattering (SERS) tags.

Figure 17. Schematic of photoelectrochemical detection of proteins. Affinity reagent immobilized on the surface binds to the target protein, resulting in steric hindrance that decreases the intensity of the photocurrent. Reprinted with permission from ref 265. Copyright 2015 American Chemical Society. Q


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single molecules.275−278 Additionally, multiplexed detection for five protein analytes has been demonstrated using arrays of microring resonators.279 Additional information on WGM is provided in refs 280 and 281. 7.1.4. Surface-Enhanced Raman Scattering (SERS) Tags. A molecule in a sample can be identified based on its Raman spectra. Raman spectroscopy on its own is not sufficient to detect protein molecules in most cases because Raman signals are weak. Therefore, surface-enhanced Raman scattering (SERS) is often used in which a metal surface, generally composed of gold or silver, enhances the Raman spectral lines. SERS tags, metal nanoparticles composed of a Raman reporter that has strong Raman scattering, a protective shell, and an affinity reagent, have been used to enhance the sensitivity of Raman-based protein assays. SERS tags have several advantages including high sensitivity,282 spectral lines with narrow line widths, and multiplexing capabilities. Furthermore, SERS tags have high stability and do not photobleach. SERS tags can be detected with NIR lasers to minimize background autofluorescence arising from components in a biological sample.283 A multiplexed sandwich assay using SERS tags as labels is shown in Figure 19.284 Different approaches have been developed for protein detection using SERS.285−287 Several excellent reviews have been written on use of SERS tags and bioassays.288−291

based signal detection could have potentially high sensitivity due to reduced background.263−266 Therefore, even if biological molecules in the sample are excited, they will likely not be detected. There are two essential components of PEC assays for protein detection: an affinity reagent that is near the signal transducer and a photoactive species. Typical molecules used for PEC signal generation are organic or inorganic semiconductors, quantum dots,267,268 or secondary enzyme labels, such as glucose oxidase, horseradish peroxidase, alkaline phosphatase, and acetylcholine esterase, that can produce photocurrent due to catalysis. Development of PEC-based assays for protein detection has gained momentum in recent years. Two general approaches have been used. The first is label-free detection in which binding of the target protein to an affinity reagent results in steric hindrance that decreases the intensity of the photocurrent.269,270 The second approach is based on a typical sandwich-assay format in which an affinity reagent binds to the target protein, and subsequent labeling with a PEC active species results in measurable photocurrent. High-sensitivity PEC protein assays with detection limits in the subpg/mL range have been developed.271 Recently, multiplexed protein detection methods based on an antibody array or two different enzyme labels that produce distinguishable PEC signals were developed.272−274 PEC assays are an emerging class of protein assays due to their potential for high sensitivity and relative simplicity. 7.1.3. Optical Ring Resonators. In an optical ring resonator, light undergoes total internal reflection along a curved boundary in a structure known as an optical microcavity, resulting in light propagation in circulating waveguide modes or whispering gallery modes (WGMs). The sensing layer is modified with affinity reagents, and when the target protein binds there is a shift in the resonance wavelength. Figure 18 shows a schematic of WGM for protein detection. WGM can have high sensitivity capable of detecting

7.2. Miniaturization

Miniaturized assays are advantageous due to size compatibility between the target protein and the assay format. Major advantages of these systems are low volume requirements and enhanced sensitivity. In this section, we discuss miniaturized approaches for protein detection including mechanical detection using microcantilever arrays and ultrasmall containers. 7.2.1. Microcantilever-Based Assays. Advances in micro- and nanofabrication technologies have enabled development of miniaturized mechanical-based assays for protein detection in which binding of a protein analyte to an affinity reagent leads to a detectable change in mass.292,293 Mechanical assays have several advantages for protein detection. They can achieve yoctogram resolution,294 with single-protein molecule detection,295 are amenable to multiplexing, are low cost, and can be used for point of care applications.296 Additionally, they are available in a label-free assay format but can also be integrated with secondary labeled affinity reagents to enhance the signal. Recently, microcanteliver assays have been used for sensitive and multiplexed protein detection based on mechanical signal transduction.297−300 In such assays, an affinity reagent immobilized on the surface of the microcantilever specifically binds to the target protein, resulting in a shift of the resonance frequency or lateral sheer stress that leads to bending of the microcantilever.301 The resonance frequency and bending can be measured using piezoresistivity or optical deflection of a light beam at the surface of the microcantilever. A scheme is shown in Figure 20.302 Microcantilevers have several advantages for protein detection One advantage of microcantilever arrays is their ability for multiplexed detection (Figure 20B).303−305 Additionally, microcantilever assays have a fast response time and low cost. They are also amenable to microfabrication technologies and integration into portable point of care devices. Microcantilevers can be used for highly sensitive and

Figure 18. Whispering gallery mode (WGM) sensor for protein detection. (a) Resonance shift is associated with protein binding. (b) WGM in a dielectric sphere. Light circumnavigates the glass sphere. Antibodies are immobilized onto the surface, and binding of the protein results in a shift of the resonance wavelength. (c) Singleprotein molecule binding events are theorized to appear as steps in the wavelength shift. Reprinted with permission from ref 275. Copyright 2008 Nature Publishing Group. R


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Figure 19. Schematic of multiplex protein sandwich assay on a porous hydrogel bead using SERS nanotags. Porous hydrogel beads are modified with capture antibodies, which bind to the target protein. Target protein is then labeled with SERS nanotags which are functionalized with a detection antibody. Raman dyes are embedded in the Au core and the Ag shell. Two different SERS nanotags have been used for multiplexed detection. Reprinted with permission from ref 284. Copyright 2018 American Chemical Society.

Figure 20. Microcantilever for protein detection. (a) Affinity reagents are immobilized onto the surface of the microcantilever. Antifouling molecules are coated onto the bottom surface of the microcantilevers to reduce nonspecific binding. Target protein then binds to the affinity reagent. Secondary gold−nanoparticle functionalized with antibodies binds to the protein. (b) SEM of the microcantilevers. (c) Optical beam deflection for measuring microcantilever vibration. (d) Mass loading onto the microcantilever results in a shift in resonance frequency. Reprinted with permission from ref 302. Copyright 2014 Nature Publishing Group.

quantitative detection of proteins.306−308 Improvements in sensitivity using secondary antibodies tagged with nanoparticles, which increase the mass response of the microcantilever assays, have resulted in sensitivity in the low fg/mL range.302 An important consideration is nonspecific binding at the bottom surface of the microcantilever. Surface passivation can be used to overcome this limitation.309 7.2.2. Ultrasmall Containers. Miniaturized systems are favorable for protein detection due to the compatibility between the target protein molecule and the size of the

container. Protein molecules are a few nanometers in size and a single cell about a few micrometers in size. Analysis performed in a test tube, in which a single protein molecule or a single cell is present in a large volume, will likely result in inability to locate, capture, and detect the protein molecule or cell. Confinement of molecules into ultrasmall containers enables detection of proteins at the single-molecule and single-cell level. Several approaches using ultrasmall volume containers for protein analysis have been developed. One example is the S


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Figure 21. Single-molecule enzyme analysis using a microwell array. (a) Enzyme F0F1 ATPase pumps protons from the outside to the inside of the well by hydrolyzing ATP. Fluorescent pH indicator is present inside the well and monitors changes in pH. (b) Fluorescence images after addition of ATP at 0 and 6000 s. Right panel (blue) shows the difference in intensity between the two time points. Reprinted with permission from ref 311. Copyright 2014 Nature Publishing Group.

single-molecule array (Simoa) immunoassay, which was discussed in section 6.2.2. In this technique, the fluorescent product of the enzyme−substrate reaction is confined to a small volume of approximately 50 fL. Another example is the SlipChip immunoassay in which a bead-based immunoassay is performed in nanoliter volumes.310 Finally, ultrasmall containers can also be used to measure the enzymatic activity of proteins. For example, this approach was used to study the transporter activity of F0F1 ATPase, which pumps protons across a membrane by hydrolyzing ATP (Figure 21).311 In this method, a fluorescent pH indicator is first added to the wells on a microwell array. The microwell array is then coated with a lipid bilayer, which mimics the lipid bilayer of a cell. Liposomes embedded with zero, one, or two molecules of the enzyme F0F1 are then introduced onto the array, and some fuse with the lipid bilayer on the microwells. ATP is then added to initiate the proton-pumping enzymatic activity of F0F1 ATPase across the lipid bilayer. Fluorescent images are then obtained to detect single-molecule activity. Another example is the Nanopore technology, which can be used to detect singleprotein molecules.312 In this method, changes in ionic current when the protein moves into the pore are monitored. Nanopores have been used to distinguish between different phosphorylation states. Ultrasmall containers have also been used for single-cell analysis. One application is detection of cytokine release from single cells using an array of wells in which only one cell can fit per well.313 Using this approach, highly multiplexed detection of secreted proteins from single cells was developed (Figure 22).314,315 In this method, single cells are first isolated inside individual microchambers. Fifteen capture antibodies specific to different cytokines are coated onto the surface of the chamber at known and predefined positions. Following a 12− 24 h incubation, the capture antibodies bind to the secreted cytokines. The barcoded chamber is then removed, and a sandwich assay is performed by using three different optical labels. Using a combination of spatial and spectral encoding, a 42-plex assay was developed. A second approach based on isolating single cells inside microwells is the single-cell Western blot, which was previously described in section 6.1.4. Another method used to quantify proteins and mRNA levels simultaneously in single mammalian cells is based on a digital proximity ligation assay (PLA) for protein quantification and

Figure 22. Highly multiplexed detection of secreted proteins from single cells. Single cells are first captured inside a microwell array. Following incubation, secreted cytokines are captured onto capture antibodies. Barcoded slide is then removed, and a sandwich assay is performed. By using three optical encodings and 15 spatial encodings, a 45-plex assay is possible. Modified and reprinted with permission from ref 315. Copyright 2015 National Academy of Sciences.

droplet digital PCR (ddPCR) for mRNA quantification (Figure 23).164 Single cells are isolated and then lysed, and the cell lysate is split into two separate samples. For digital PLA, the cell lysate sample is incubated with oligonucleotidebound antibodies. The antibodies bind to the target protein, bringing the oligonucleotides near each other. A connector oligonucleotide is added that hybridizes to the probes to form a complex consisting of the target protein analyte, the probes, and connector oligonucleotide. Ligation is performed, forming double-stranded DNA (dsDNA) followed by proteolytic digestion, resulting in a solution containing the dsDNA. Using a commercially available device for ddPCR (BioRad), the solution containing the dsDNA is diluted and emulsified to T


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Figure 23. Digital Droplet PCR for quantification of proteins and mRNA in single cells. Single cells are isolated and then lysed, and the cell lysate is split into two separate samples. For protein detection, PLA is performed followed by proteolytic digestion, resulting in a solution containing dsDNA. Solution containing the dsDNA is diluted and emulsified to form about 20 000 nL droplets, such that each droplet contains either 1 or 0 dsDNA molecules. Each dsDNA molecule is then amplified by PCR, and digital counting is performed by measuring the fluorescence using a droplet reader. For mRNA detection, digital droplet PCR is performed. In this manner, both protein and mRNA from a single cell can be measured. Reprinted with permission from ref 164. Copyright 2016 Cell Press.

protein measurements. Ultrasensitive methods such as the Meso Scale Discovery assay (MSD) and single-molecule arrays (Simoa) are being used to detect previously unmeasurable levels of proteins. Recent advances from the literature, such as novel optical labels and assay miniaturization, enable protein detection on the single-molecule and single-cell level. These assays will undoubtedly lead to interesting new discoveries in biology as well as new clinical diagnostics applications.

form about 20 000 nanoliter droplets, such that each droplet contains either one or zero dsDNA molecules. Each dsDNA molecule is then amplified by PCR, and digital counting is performed by measuring the fluorescence using a droplet reader. Digital PLA using ddPCR is combined with RT-ddPCR for simultaneous quantification of both protein and mRNA from a single cell. For protein quantification, femtomolar sensitivity corresponding to approximately 10 000 molecules was achieved with a dynamic range of about 3 orders of magnitude. For mRNA quantification, a limit of detection of approximately 52 mRNAs per cell was achieved.

AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected].

8. CONCLUSION AND FUTURE DIRECTIONS In this review, we discussed the wide variety of approaches for protein detection. Several parameters, including sensitivity, specificity, dynamic range, multiplexing capabilities, reproducibility, ease of use, and cost, are important in protein detection techniques. A major analytical challenge with current protein detection techniques is measuring multiple proteins that are present in complex biological samples with high sensitivity and a wide dynamic range. Strategies to overcome these challenges have been developed; however, there is still a trade-off between high-sensitivity measurements and highly multiplexed measurements of proteins. As a result, it is often necessary to decide whether it is more advantageous to detect several specific proteins using a candidate-based, hypothesisdriven approach or to detect as many proteins as possible using an unbiased, high-throughput approach that is less sensitive. A major challenge in ultrasensitive and highly multiplexed protein detection is nonspecific binding between biomolecules in the sample, assay reagents, and surfaces. Nonspecific binding leads to background signals that obscure target protein binding. Surface passivation methods that reduce nonspecific binding as well as development of high-affinity reagents that do not cross-react will enhance the performance of protein detection methods. In addition, low-cost protein detection methods for applications in developing countries or point of care clinical applications are still necessary. Nevertheless, major advances in protein detection methods have been developed in recent years. One example is the SOMAscan assay, which is capable of highly multiplexed

ORCID

Limor Cohen: 0000-0003-1448-0925 David R. Walt: 0000-0002-5524-7348 Notes

The authors declare the following competing financial interest(s): David R. Walt is the scientific founder and a board member of Quanterix Corporation. All other authors declare no competing financial interest. Biographies David R. Walt is Professor of Pathology at Harvard Medical School and Brigham and Women’s Hospital, a Core Faculty Member of the Wyss Institute at Harvard University, and a Howard Hughes Medical Institute. He is the Scientific Founder of multiple life sciences companies. He has received numerous national and international awards for his work in the field of optical microwell arrays and single molecules. In addition to his lab’s fundamental work on single molecules, his lab develops new diagnostics technologies and applies them to pressing clinical problems. He is a member of the National Academy of Engineering, the National Academy of Medicine, and a Fellow of the American Academy of Arts and Sciences, the American Institute for Medical and Biological Engineering, and the National Academy of Inventors. Limor Cohen is a Ph.D. candidate in the Department of Chemical Biology at Harvard University. She received her B.A. degree in Chemistry from New York University in 2012. Her research interests U


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