Painting protein misfolding in the cellin real time with an atomic-scale brushSilvia Cavagnero and Lisa M. Jungbauer

Department of Chemistry, University of Wisconsin-Madison, 1101 University Ave, Madison, WI 53706, USA

The direct observation of specific biochemical events in

living cells is now possible as a result of combined

advances in molecular biology and fluorescence micro-

scopy. By genetically encoding the source of a unique

spectroscopic signal, target proteins can be selectively

detected within the complex cellular environment, with

limited interference from background signals. A recent

study takes advantage of arsenical reagent-based

methodologies to monitor in vivo protein misfolding

and inclusion body formation in real time. This approach

promises to yield important information on the kinetics

of aggregate formation in living cells and its relation

to the time-course of protein expression and post-

translational processing. The ability to follow protein

self-association in real time accurately from its early

stages is unique to this method, and has far-reaching

implications for both biotechnology and misfolding-

based disease.

Introduction

Cell biology has recently witnessed spectacular advancesas a result of novel technologies that enable the fate ofindividual proteins to be followed with high specificity inliving cells. Macromolecular interactions are often moni-tored by the in vivo yeast two-hybrid system screeningassay and related variants [1]. This methodology indirectlyreports on target protein interactions through the restoredfunction of reconstituted reporter enzymes. A more directand quantitative strategy to map both macromolecularlocalization and protein–protein interactions in the cellinvolves the combination of fluorescence microscopyimaging (or spectroscopy) and the genetic incorporationof fluorogenic amino acid sequences into a target protein.Fusion constructs combining a highly fluorogenic proteinreporter with the sequence of the biomolecule of interestare commonly used. The reporter moiety describes theintracellular properties and spatial distribution of theprotein of interest with high selectivity [2,3]. The mostpopular amino acid sequence used as a source of intensefluorescence emission is the green fluorescent protein(GFP) from the jellyfish Aequorea victoria and itsnumerous variants [4,5]. Related proteins from otherorganisms with complementary spectroscopic propertieshave also been employed [6]. Cellular localization ofindividual proteins is monitored by simple fluorescenceemission, whereas colocalization of multiple proteins and

Corresponding author: Cavagnero, S. (cavagnero@chem.wisc.edu).Available online 28 January 2005

www.sciencedirect.com 0167-7799/$ - see front matter Q 2005 Elsevier Ltd. All rights reserved

protein–protein interactions is typically monitored byfluorescence resonance energy transfer (FRET) variantsof this approach.

In general, the reporter protein-based approach isrobust and widely used. However, it has the disadvantagethat the large size of the reporter protein (e.g. GFP has238 amino acids) might perturb the intracellular proper-ties of the protein of interest. Although this has notgenerally been a problem, investigations involving inter-molecular interactions with large surface contacts, such asextensive protein aggregation, must take this issue intoaccount. For example, it has been reported that proteinaggregation alters the fluorescence of a covalently boundGFP reporter by preventing its proper folding [7].

The in vivo post-translational maturation of GFPand its analogs (involving an intramolecular oxidativecyclization) is very slow and the apparent half-life for thegeneration of GFP fluorescence ranges between 30 and90 minutes [8]. Faster maturing proteins from jellyfishand other organisms are under development [9], includingenhanced versions of yellow fluorescent protein (YFP) andGFP, denoted as Venus and T-sapphire, which acquirefluorescence with a remarkable apparent half-life of only1.5 and 5 minutes, respectively [10,11]. Therefore, kineticstudies of intracellular processes occurring on the minutetimescale or faster are not possible unless a Venus-fusionprotein approach is used. However, the bulky size of theadded fluorophore is not ideal in kinetic and mechanisticstudies involving extensive intermolecular contacts.These investigations are better served by an alternativefast-maturing system using a smaller fluorophore.

FlAsHing light into specific biochemical processes

Tsien and coworkers have developed a clever arsenic–fluorescein-based approach employing an engineeredreceptor–ligand pair that becomes fluorescent upon com-plexation [12]. This method enables the specific fluor-escent labeling of any desired protein expressed insideliving cells with only moderate background signal.Figure 1 schematically illustrates the method [13]. Briefly,a short tetracysteine motif (Cys-Cys-Xaa-Xaa-Cys-Cys,where Xaa denotes any generic amino acid, preferablyPro-Gly [14]) is engineered into the target protein. Twoproperly positioned arsenic atoms in a custom-madebiarsenical reagent selectively coordinate the tetracys-teine motif. The fluorophore-derivatized biarsenicalligand is known as FlAsH-EDT2 (fluorescein arsenicalhelix binder, bis-ethanedithiol adduct). This compound is

Review TRENDS in Biotechnology Vol.23 No.3 March 2005

. doi:10.1016/j.tibtech.2005.01.008

+ 2

Cys

Cys

+

Cys

Cys

Cys

S

S

SS

SHSH

SHSH

FlAsH-EDT2

FlAsH

O OHO

COOH

As AsSS SS

O OHO

COOH

As

HS SH

As

Cys

Cys

Cys

TRENDS in Biotechnology

Figure 1. Site-specific protein labeling by the FlAsH fluorescent arsenical reagent. The primary sequence of the protein of interest is genetically engineered by inserting the

CCXXCC sequence (CZcysteine and XZany other amino acid) at the amino- (or carboxy-) terminus or an exposed surface loop. Upon addition of the fluorescein arsenical

hairpin binder (FlAsHe-EDT2, green star) labeling reagent, the engineered cysteine residues coordinate the arsenic, displacing ethane dithiol (EDT) and converting the dye to

its fluorescent state. The figure displays FlAsH labeling of the cellular retinoic acid-binding protein I (CRABP I) carried out in thework by Ignatova and Gierasch [36], discussed

in this review. The protein image was generated with the software Protein Explorer from the Protein Data Bank (PDB) file 1CBI [44].

Review TRENDS in Biotechnology Vol.23 No.3 March 2005158

generally cell membrane-permeable and it can be conveni-ently preloaded into cells before expression of the targetprotein. Remarkably, the ligand is virtually non-fluorescentuntil bound to its tetracysteine receptor. Binding takesplace rapidly (with a half-life of w1.5 minutes). Uponaddition of the FlAsH reagent, some undesired back-ground fluorescence is always generated in vivo. This isprimarily due to the presence of endogenous cellularproteins displaying a weak affinity for FlAsH. Thesegenerally bear one or more cysteines at variable positions.The background signal is minimized by treating the cellswith moderate amounts of ethane dithiol (EDT), thusensuring that excess FlAsH is more likely to undergoexchange with EDT, rather than binding to endogenouscysteine-containing proteins*.

Since the original implementation, other FlAsH deriva-tives with complementary spectroscopic properties havebeen designed [14,15], including an analog with anenvironmentally sensitive fluorophore on the biarsenicalligand for in vivo imaging of protein-conformationalchanges [16]. Widespread applications of the FlAsHtechnology include selective labeling of cell-surface pro-teins, affinity purification [17] and direct in-gel detectionof expressed proteins [14]. One of the modified biarsenicalanalogs, ReAsH, is both fluorescent and suitable fordetection by electron microscopy [14], providing a power-ful tool for correlating optical and electron microscopyimages. FlAsH–tetracysteine labeling has been used toinvestigate several in vivo biochemical events. Theseinclude gap junction assembly and turnover [18], proteinexpression levels [19], protein function probed by selective

* The biarsenical FlAsH reagent is currently marketed by Invitrogen (http://www.invitrogen.com) under the Lumioe trade name.

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in situ photoinactivation [20,21] and trafficking of anEbola virus matrix protein in live cells [22]. In the specificcase of in vivo protein misfolding kinetics, FlAsH-typelabeling offers competitive advantages over GFP and otheravailable fluorescence-based methods (see Table 1 for acomprehensive overview). The fluorogenic moiety is smalland it is rapidly co- or post-translationally incorporated,provided that cells are preloaded with FlAsH-EDT2 beforetarget protein expression.

From inclusion bodies to Lewy bodies: protein

misfolding in biotechnology and disease

Protein misfolding is a highly undesirable process in thecontext of living cells. In biotechnology and basic research,overexpression of target genes often generates macro-scopic solid deposits known as inclusion bodies. Theseusually require laborious procedures for their conversionto soluble native-like active protein [23]. The generation ofheavily misfolded proteinaceous aggregates is directlylinked to the pathology of several human diseases. Forexample, nuclear inclusions, amyloid fibrils or Lewybodies are generated intracellularly in the case ofHuntington’s, Alzheimer’s and Parkinson’s neurodegene-rative disorders [24], respectively. In vitro studies moni-toring the presence and formation of self-associatedspecies have been instrumental in revealing fundamentalaspects of this process. However, it is only through in vivoinvestigations that the actual cellular localization andbiologically significant self-association mechanisms can beelucidated.

Several studies have been successful at mappingthe distribution of macroscopic protein aggregates in thecell by imaging techniques, mostly based on fluorescence

Table 1. Available methods for tracking macromolecular behavior in vivo by genetically encoded site-specific fluorescence labeling

Review TRENDS in Biotechnology Vol.23 No.3 March 2005 159

microscopy. For example, the senile plaques of Alzheimer’sdisease have been studied by multiphoton microscopyupon thioflavin S staining in live mice [25]. Details onintermolecular interactions among Parkinson’s disease-related Lewy bodies have been obtained by in vivodetection of aggregates tagged by fluorophore-labeledsecondary antibodies in brain tissues [26]. Rajan et al.have used GFP-based FRET combined with deconvolutionmicroscopy to monitor the specificity of inclusion bodyformation in intact eukaryotic cells [27]. GFP-basedstudies monitoring fluorescence recovery after photo-bleaching (FRAP), fluorescence loss in photobleaching(FLIP) and FRET have characterized the diffusivemotionsand interaction among polyglutamine–huntington protein(Htt) aggregates and other glutamine-rich proteins or theHsp70 chaperone in vivo [28].

The above work has significantly contributed to ourunderstanding of the distribution of aggregates in vivo.However, the fundamental reasons behind intracellularself-association are far from being understood, and morestudies are urgently needed to explore the mechanisticaspects of this issue in a biologically relevant context.A first important step in this direction relies on improvedmethods to monitor in vivo aggregate formation in realtime, including the early events that take place beforemacroscopic fibrils or other large complexes are generated.Previous work was based primarily on studies employingGFP as an aggregation reporter. Specifically, the time-course of aggregate appearance was monitored for variousversions of the Htt-bearing C-terminal ‘enhanced’ greenfluorescence protein (EGFP) tags [29]. The kinetics ofpolyglutamine aggregate formation has also been moni-tored in COS-7 cells upon expression of GFP-containingconstructs reporting on the self-association of

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polyglutamine-containing atrophin-1 protein [30], whosemisfolding is linked to dentatorubral-pallidoluysian atro-phy (DRPLA), a severe neurodegenerative disorder. Theabove GFP-based reporters bear the disadvantage ofacquiring fluorescence very slowly after biosynthesis,thereby masking the aggregation stages taking placeduring the initial one or two hours. Although earlyaggregation events might be captured with Venus- orT-Sapphire-like GFP-derivatives, the large size of thetag might interfere with the self-association process,and no studies of this type have been published todate. An alternative FlAsH-based method is described inthe next section.

Watching protein misfolding in real time

In order to properly understand the mechanisms ofprotein misfolding in the cell and its implications forbiotechnology and disease, it is necessary to follow theprocess from its initial steps. Early intermediates andsmall transient aggregates are believed to play importantroles in the metabolism of the cell and to affect theultimate outcome of self-assembly processes [31,32].Therefore, observation of the protein aggregation time-course within a cell is best achieved by probes capable oftracking the full evolutionary history of a protein. Thisstarts from the generation of ribosome-bound nascentpolypeptides up to full-length chains and inclusion bodies,as illustrated in Figure 2.

The ability to follow aggregation from its early stagesposes the most stringent technical challenges. As seenabove, approaches based on GFP, its improvedmutants andother GFP analogs from different organisms suffer fromundesirable drawbacks. Alternative approaches basedon radioactive labeling [33,34] or antibody tagging [35]

TRENDS in Biotechnology

Ribosome-boundnascent chains

Full-lengthpolypeptides

Inclusionbodies

.

.

....

.

.

.

.

.

.

mRNA

PolypeptideRibosome

Figure 2. Schematic representation of in vivo inclusion body formation. Ribosomal

protein biosynthesis leads to accumulation of full-length polypeptide chains. In

some cases, the newly synthesized polypeptides do not properly fold to their native

state, and form macroscopic aggregates known as inclusion bodies. Co- or post-

translational incorporation of the FlAsH fluorescence reagent (green star) enables

monitoring of the time course of in vivo protein synthesis and the subsequent

inclusion body formation.

Review TRENDS in Biotechnology Vol.23 No.3 March 2005160

might perturb the cell’s equilibrium because they involvethe physical separation of cellular components after self-association has taken place.

Ignatova and Gierasch have taken advantage of thefast incorporation of the biarsenical FlAsH reagent into anengineered tetracysteine motif (Figure 1) to monitor thetime-course of protein misfolding in Escherichia coli cells[36]. Two time points are shown in Figure 3. FlAsH-EDT2

is only fluorescent upon binding to the tetracysteine motif,and unincorporated FlAsH makes a negligible contribu-tion to background fluorescence. The FlAsH–tetracysteineacceptor sequence can, in principle, be appended to eitherthe amino- or carboxy-terminus or inserted anywhere inthe interior of the sequence of a protein, provided thatno significant structural perturbations are introduced.

Figure 3. In vivo visualization of inclusion body formation in real time by

fluorescence microscopy. Images of agarose-immobilized cell suspensions

obtained 60 (left) and 240 (right) minutes after isopropyl-beta-D-thiogalactopyrano-

side (IPTG)-promoted P39A tetra-Cys CRABP I expression. The inclusion body

fluorescence is due to FlAsH-EDT2 probe incorporation. After 60minutes, very small

fluorescent aggregates are visible for only a few cells. By 240 minutes, the majority

of the cells have accumulated a large number of highly fluorescent aggregates.

Reproduced, with permission, fromRef. [36]. Copyright (2003) National Academy of

Sciences, USA.

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Insertion into a-helical secondary structure motifs hasproven to be quite effective [12]. Ignatova and Gieraschshowed that flexible internal loops are also good insertionsites for the FlAsH binding motif [36].

Given that the FlAsH-EDT2 dye is only moderately cellpermeable, intact cells had to be pretreated with moderateamounts of lysozyme and then washed, before isopropylb-D-thiogalactoside (IPTG) induction. This procedurepartially disrupts the peptidoglycan layer and it facili-tates the intracellular incorporation of the biarsenicalFlAsH-EDT2 dye. The E. coli outer cell membranerecovers its original nature within one generation, oncelysozyme has been removed. Cells pretreated by thismethod appear morphologically intact by fluorescencemicroscopy (Figure 3) and are as viable as untreatedcontrols. The cellular retinoic acid binding protein I(CRABP I) is mostly composed of b-sheets and it usuallyexpresses in soluble form. However, its P39A mutantforms inclusion bodies. This protein pair is therefore agood system for monitoring variations in the FlAsHfluorescence reporter as a function of protein aggregation.Self-association and inclusion body formation is monitoredby both fluorescence microscopy imaging and fluorescenceemission spectroscopy in bulk cell suspensions. Assumingthat the biarsenical dye has been internalized at the timeof protein synthesis initiation, and considering the fasttime scale of FlAsH binding, the aggregation time-courseis followed from its very early steps. The P39A mutantaggregates fast and to amuch larger extent than wild-typeCRABP I. Interestingly, this in vivo overexpressed mutantcan be refolded to a native-like monomeric conformationunder diluted in vitro conditions. However, this processoccurs significantly more slowly than for the wild-typeprotein [37], suggesting that the extra time intrinsicallyrequired for P39A folding might enable kineticallycompeting misfolding processes to take over and dominatethe conformational search in vivo.

A key issue in these studies is that the quantum yieldof FlAsH-labeled CRABP I fluorescence depends on itsconformation and chemical environment. For example,urea-unfolded FlAsH-labeled CRABP I has a significantlyhigher fluorescence than its native state. In whole cells,the fluorescence signal increases upon inclusion bodyformation. This suggests the possible presence of rela-tively unfolded pre-inclusion bodies or mature aggregates.This aspect of the work will undoubtedly be enriched byadditional future investigations taking the quantumyields of individual species into account. The approachby Ignatova and Gierasch is simple and powerful, andholds promise to become routinely employed to follow pro-tein self-association in vivo, starting from its early stages.

Challenges and future directions

Although the FlAsH technology holds great promise as atool for the study of protein self-association kinetics in thecell, several potential drawbacks might somewhat limitits general applicability. For example, FlAsH-mediatedfluorescence lacks a well-defined characteristic spectro-scopic signature for conformationally distinct populations(i.e. the unfolded, folded and aggregated states), in thatno significant emission frequency shifts are typically

Review TRENDS in Biotechnology Vol.23 No.3 March 2005 161

observed for the different states. Fluorescence quantumyields vary in a somewhat unpredictable fashion. There-fore, careful parallel in vitro and in vivo control experi-ments are required to characterize the fluorescencecontributions due to each state and the correlationbetween fluorescence response and aggregation-drivenmisfolding. Deconvolution of the observed in vivo fluor-escence signal is straightforward only in the presence of alimited number of species bearing distinctly differentfluorescence quantum yields.

Efficient FlAsH labeling requires the cysteine thiols tobe present in reduced form, limiting the applicability ofthis method to reducing intracellular environments[13,14]. Therefore, FlAsH labeling of proteins targeted toorganelles such as the endoplasmic reticulum and secre-tory vesicles, which support oxidizing conditions, is likelyto be inefficient. In the presence of competing cysteineoxidation, undesired inter- or intramolecular disulfidebridges are formed. However, artificial alteration of localredox properties can be successfully applied to overcomethe above complication, as demonstrated for the labelingof a vesicle-associatedmembrane protein inHeLa cells [14].

Another potential limitation of the FlAsH labelingsystem is the presence of background signal. Three typesof background fluorescence are possible: nonspecificlabeling of endogenous cysteines (discussed previously),nonspecific FlAsH binding to hydrophobic sites and cellu-lar autofluorescence. In the majority of FlAsH-basedstudies, an acceptable signal-to-noise ratio is obtained byincluding EDT in the FlAsH labeling solution. Thisminimizes fluorescence due to nonspecific cysteine thiollabeling. However, in several cell lines, residual highbackground persists (e.g. in HEK 293 and CHO cells [38]).Thiol-independent background signal (due to FlAsHbinding to hydrophobic protein and organelle pockets, asin ECV 304, HEK 293, CHO, 3T3 and 3T6 cells [13]) can beattenuated by the co-delivery of uncharged membrane-permeant dyes such as Disperse Blue 3 [13]. Althoughcellular autofluorescence and light scattering have notbeen cited as significant problems for the detection ofFlAsH fluorescence, a red-shifted FlAsH analog such asReAsH (608 nm emission maximum wavelength) reducesthe potential for scattering-related background signal.

As discussed above, FlAsH delivery into E. coli isfacilitated by increasing the bacterial cell wall perme-ability by lysozyme pretreatment [36]. However, FlAsH isreadily membrane-permeable in a variety of eukaryoticcells [13], including Drosophila larvae [21], HeLa cells[12,13,18,20], Jurkat lymphocytes [12], skeletal muscle(dysgenic myotubes) [38], 293Tcells [13,22] and yeast cells[39]. Although FlAsH treatment is typically carried out for30 minutes up to several hours to enable complete cellpermeation, dye preloading into cells reduces the experi-mental dead time and it enables fast labeling, to followthe early kinetics of intracellular events [36]. A creativeexperiment employing pulse-chase labeling of gap junc-tion proteins with FlAsH and ReAsH dyes in HeLa cellssuggests that the preloading strategy is also readilyapplicable to eukaryotic cells because protein expressioncontinues effectively (for up to 8 hours) after initial FlAsHlabeling [18]. FlAsH labeling has been extended to live

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dissected Drosophila larvae [21]. However, it is of limitedutility in whole live organisms because the need fornoninvasive dye delivery and removal of excess label priorto imaging poses serious challenges.

FlAsH-based technology promises to provide muchneeded insights into the kinetics of in vivo protein self-association, starting from its early stages. This is signi-ficant for both biotechnology and medicine because earlyaggregation stages are likely to be important in directingthe fate of the newly synthesized cellular proteins. Sortingout and controlling the pathways leading to irreversibleaggregation, degradation or rescuing by the many helpersavailable inside the cell (including chaperones) is one ofthe most exciting outstanding challenges in cell biology.

AcknowledgementsS.C.’s group is funded, in part, by the National Science Foundation(MCB-0215368) and by the National Institutes of Health(1R01GM068535-01A1). L.M.J. was supported by a Predoctoral TrainingGrant in Molecular Biophysics (5 T32 GM-09283).

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