Superresolution imaging of targeted proteins in fixed and living cells using photoactivatable organic fluorophores

Article abstract of DOI:10.1021/ja1044192

Superresolution imaging techniques based on sequential imaging of sparse subsets of single molecules require fluorophores whose emission can be photoactivated or photoswitched. Because typical organic fluorophores can emit significantly more photons than

Full text of DOI:10.1021/ja1044192

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J Am Chem Soc. Author manuscript; available in PMC 2011 November 3.
Published in final edited form as:
J Am Chem Soc. 2010 November 3; 132(43): 15099–15101. doi:10.1021/ja1044192.
Superresolution Imaging of Targeted Proteins in Fixed and
Living Cells Using Photoactivatable Organic Fluorophores
1
1
1
2
2
Hsiao-lu D. Lee , Samuel J. Lord , Shigeki Iwanaga , Ke Zhan , Hexin Xie , Jarrod C.
4
4
3
3
3
4
Williams , Hui Wang , Grant R. Bowman , Erin D. Goley , Lucy Shapiro , Robert J. Twieg ,
2
1
Jianghong Rao , and W. E. Moerner
1Department of Chemistry, Stanford University, Stanford, California 94305
2Department of Radiology, Stanford University, Stanford, California 94305
3Department of Developmental Biology, Stanford University, Stanford, California 94305
4Department of Chemistry, Kent State University, Kent, Ohio 44244
Abstract
Superresolution imaging techniques based on sequential imaging of sparse subsets of single
molecules require fluorophores whose emission can be photoactivated or photoswitched. Because
typical organic fluorophores can emit significantly more photons than average fluorescent
proteins, organic fluorophores have a potential advantage in superresolution imaging schemes, but
targeting to specific cellular proteins must be provided. We report the design and application of
HaloTag-based target-specific azido DCDHFs, a class of photoactivatable push–pull fluorogens
which produce bright fluorescent labels suitable for single-molecule superresolution imaging in
live bacterial and fixed mammalian cells.
Recently, sequential imaging of sparse subsets of photoactivatable/photoswitchable single-
molecule fluorophores has enabled optical imaging beyond the diffraction limit (DL),
1–3
providing insight into the sub-diffraction world (e.g. PALM, FPALM, STORM).
These
single-molecule superresolution (SR) techniques have provided the impetus for development
of new controllable fluorophores with large numbers of emitted photons N, because the
4
5
achievable resolution scales as
. Most previous SR experiments in living cells have
6–9
used photocontrollable fluorescent proteins.
However, despite having the advantage of
being target-specific, fluorescent proteins on average provide 10-fold fewer photons before
10,11
photobleaching than good organic fluorophores.
Small organic fluorophores have the
additional benefit of synthetic design flexibility for tuning target specificity, spectral
wavelength, solubility, and other desired properties. Therefore, targeted bright organic
wmoerner@stanford.edu .
Ref. 20
. This material is
Supporting Information Available: Procedures, chemical synthesis, analysis, additional figures, and complete
available free of charge via the Internet at http://pubs.acs.org.

Lee et al.
Page 2
fluorophores which are compatible with the live-cell environment and which can be turned
on and/or off would be advantageous. While targeted fluorogens which activate via esterase-
12
mediated cleavage have been previously demonstrated, optical control of the emitting
13
concentration is necessary for SR.
Many biological processes can be interrupted by fixing (i.e. killing) cells, which is required
16,17
for the immunostaining used in most SR experiments that rely on organic fluorophores.
While fixation can be useful, there is a need for techniques and probes that can be used for
SR imaging in living cells. Single-molecule imaging in living cells using exogenous
5
fluorophores faces the dual hurdles of cell permeability and targeted labeling. There are
exceptions in the literature that did not use fixation and immunostaining. For instance,
17
Heilemann et al. imaged mRNA in living cells using oligomers labeled with small organic
18
fluorophores and Conley et al. labeled the external lysines of bacterial cells using a Cy3–
Cy5 heterodimer. Very recently, fluorophores were targeted with trimethoprim and intrinsic
19
cellular reductants enabled photoinduced blinking . These examples demonstrated some
possibilities for live-cell labeling and SR, but there is still a need for photoactivatable
organic labels for SR imaging because each molecule is localized only once, while with
blinking or photoswitching, each molecule can be localized a variable number of times.
Here we present a target-specific photoactivatable organic fluorophore for use inside living
20–22
and fixed cells, 3, based on the commercial HaloTag targeting approach.
This method
requires a genetic fusion to the HaloEnzyme (HaloEnz), which forms a covalent linkage to
the HaloTag substrate, thus labeling the protein of interest (i.e. a protein–HaloEnz–
HaloTag–fluorophore covalent unit). Specifically, we present: (i) the basic photophysical
properties of a new targeted photoactivatable probe; (ii) proof-of-principle labeling of
known structures in fixed and living mammalian cells validated by co-staining with
antibodies or co-transfection with fluorescent proteins; (iii) specific SR imaging of
microtubules in a mammalian cell with quantification of resolution enhancement; (iv)
demonstration of targeted labeling in living bacteria with diffraction-limited imaging; and
finally, (v) SR imaging of poorly understood structures inside living bacteria.
As molecules with bright emission for single-molecule imaging,
dicyanomethylenedihydrofuran (DCDHF) push–pull fluorophores emit millions of photons
15,23
before photobleaching, and can enter living cells.
Recently, we reported a
photoactivatable DCDHF fluorogen based on photocaging the fluorescence by replacing the
amine donor with a poorly-donating but photolabile azide, which can then be converted back
14
to an amine using low-intensity blue light. Azido DCDHF fluorogens exhibit high turn-on
ratios (the increase in emission from the fluorescent form compared to the dark fluorogen)
15
of 325–1270 fold, an attractive property for SR imaging. (For a detailed discussion of the
photophysical properties of the fluorogens, as well as the methods used to characterize them,
references 11
15
see
and .)
14
Compared with the original DCDHF-V-P-azide, 3 exhibits similar spectral changes upon
optical pumping of the aryl azide (Figure 1) but a higher photoconversion quantum yield
(Φ , see Table 1). Presumably the oxygen on the aryl azide stabilizes the intermediate
P
14,15,24
nitrene, making photoconversion more favorable.
The photoconversion of 3 was so
sensitive such that an additional activating blue laser was not necessary; instead, the diffuse
ambient light (e.g. the blue light emitted from a nearby computer monitor in an otherwise
dark room) was sufficient to activate sparse sets of the fluorogen. The thermal activation rate
and activation by the 594 nm imaging laser were significantly lower, as measured in
complete darkness in a covered sample (see SI). Because the fluorogen's sensitivity to
photoconversion is so high, it was possible to set the level of the ambient light such that the
bleaching rate from 594 nm pumping was similar to the activation rate. For instance, with
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Lee et al.
Page 3
room lights off and a nearby computer monitor on, we maintained a steady-state
concentration of isolated emitters. This reduced the complexity of the experiment by
17
allowing the use of only one laser. Moreover, because no blue or UV laser was necessary
for activation, photodamage to the imaging sample was greatly reduced. (The drawback to
this high sensitivity is that it increases the difficulty of preventing photoactivation before
imaging. We successfully minimized preactivation by performing all preparations in
complete darkness or under dim red lights only.)
We verified the specificity of 3 in mammalian culture. Wild-type HeLa cells and HeLa cells
transfected to express HaloEnz–α-tubulin were stained live with 3, fixed and immunostained
with Alexa488-mAb to α-tubulin. Fluorescence images after photoactivation clearly
demonstrate that 3 is only retained in the cells that expressed–HaloEnz-α-tubulin (Fig. 2A–
G). Next, live CHO cells were co-transfected with HaloEnz-α-tubulin and α-tubulin–eGFP,
and the labeling by 4 was shown to co-localize well with the eGFP labeling (Fig. 2H–I).
Most importantly, BS-C-1 cells were transfected with HaloEnz-α-tubulin, fixed, stained with
3 and washed before SR imaging by PALM. Comparing DL and SR fluorescence images
(Fig. 2J,K), the microtubule structure is clearly imaged with resolution beyond the optical
diffraction limit. After corroborating the utility of the DCDHF fluorophores for labeling
known structures in mammalian cells, we then moved our attention to cells with protein
organization that is not fully understood.
Bacteria are tiny (about a thousand bacteria could fit within one HeLa cell), and the details
of protein localization in prokaryotes are poorly understood, yet essential for function and
25
phenotype. DL imaging of labeled proteins in bacteria only produces diffuse blobs,
complicating meaningful interpretation. For these reasons, living bacterial studies benefit
11,26
greatly from SR imaging.
We used HaloTag-DCDHFs to highlight protein localization
26–29
patterns in live Caulobacter crescentus bacteria,
which demonstrates the biologically
interesting ability to divide asymmetrically. Elucidating the mechanisms of asymmetric cell
division and intracellular organization requires understanding how cytoskeletal proteins
25
27
localize through the life cycle of the cell. In this Communication, a polar protein PopZ,
28
29
and mid-plane proteins FtsZ and AmiC were expressed as HaloEnz fusions. PopZ, FtsZ,
and AmiC have distinct roles: PopZ anchors the chromosomal origin at the “swarmer” pole;
FtsZ and AmiC are recruited to the mid-plane and are components of the cell division
25
machinery. DL imaging using HaloTag targeting of the (nonphotoactivatable) fluorophore
4 shows correct PopZ localization at cell poles and FtsZ at the cellular division plane as
expected (Figure 3), confirming that this HaloTag labeling system does not significantly
interfere in phenotype.
SR images produced by photoactivation of fluorogen 3 display the expected localization
patterns, but also reveal additional detail unseen in the DL images of Figure 3. For PopZ at
the cell pole, the protein forms an asymmetric cap-like structure with a curvature that hugs
the shape of the bacterial membrane (Figure 4A–C). Also, in the case of AmiC, the protein
localizes to the cellular mid-plane, as expected (Figure 4D–E). The SR images of AmiC may
reveal a tighter organization than seen in DL microscopy, but further study is necessary
before any definitive statement can be made. In either case, these SR images provide new
detail not available in DL images.
The target-specific DCDHF single-molecule fluorogen presented here represents the first
successful installation of target-specificity to small organic photoactivatable fluorogens for
single-molecule SR imaging. Compared to existing schemes, the photoactivation of the
30
17
fluorogen does not require other additives (e.g. thiols for Cy5, or redox chemicals ) nor
activation by UV light, and thus can be used inside living cells. SR imaging has been
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directly demonstrated for fixed mammalian and live bacterial cells; additional effort to
improve washout for live mammalian cells is an important topic for future work. This and
future photoactivatable fluorogens should be helpful tools for SR imaging in the complex
environment within the living cell.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
This work was supported in part by Grant No. R01-GM086196 from the National Institute of General Medical
Sciences. We thank S. Pfeiffer for BS-C-1 cells.
REFERENCES
(1). Betzig E, Patterson GH, Sougrat R, Lindwasser OW, Olenych S, Bonifacino JS, Davidson MW,
Lippincott–Schwartz J, Hess HF. Science. 2006; 313:1642–1645. [PubMed: 16902090]
(2). Hess ST, Girirajan TPK, Mason MD. Biophys. J. 2006; 91:4258–4272. [PubMed: 16980368]
(3). Rust MJ, Bates M, Zhuang X. Nat. Methods. 2006; 3:793–795. [PubMed: 16896339]
(4). Thompson RE, Larson DR, Webb WW. Biophys. J. 2002; 82:2775–2783. [PubMed: 11964263]
(5). Lord SJ, Lee HD, Moerner WE. Anal. Chem. 2010; 82(6):2192–2203. [PubMed: 20163145]
(6). Dickson RM, Norris DJ, Tzeng YL, Moerner WE. Science. 1996; 274:966–969. [PubMed:
8875935]
(7). Ando R, Mizuno H, Miyawaki A. Science. 2004; 306:1370–1373. [PubMed: 15550670]
(8). Patterson GH, Lippincott–Schwartz J. Science. 2002; 297:1873–1877. [PubMed: 12228718]
(9). Hess ST, Gould TJ, Gudheti MV, Maas SA, Mills KD, Zimmerberg J. Proc. Nat. Acad. Sci.
(USA). 2007; 104:17370–17375. [PubMed: 17959773]
(10). Schmidt T, Kubitscheck U, Rohler D, Nienhaus U. Single Mol. 2002; 3:327.
(11). Thompson MA, Biteen JS, Lord SJ, Conley NR, Moerner WE. Methods Enzym. 2010; 475:27–
59.
(12). Watkins RW, Lavis LD, Kung VM, Los GV, Raines RT. Org. Biomol. Chem. 2009; 7:3969–
3975. [PubMed: 19763299]
(13). Maurel D, Banala S, Laroche T, Johnsson K. ACS Chem. Biol. 2010; 5:507–516. [PubMed:
20218675]
(14). Lord SJ, Conley NR, Lee HD, Samuel R, Liu N, Twieg RJ, Moerner WE. J. Am. Chem.. Soc.
2008; 130:9204–9205. [PubMed: 18572940]
(15). Lord, SJ.; Lee, HL.; Samuel, R.; Weber, R.; Liu, N.; Conley, NR.; Thompson, MA.; Twieg, RJ.;
Moerner, WE. J Phys Chem. B. 2009. ASAP. (http://dx.doi.org/10.1021/jp907080r)
(16). Folling J, Bossi M, Bock H, Medda R, Wurm CA, Hein B, Jakobs S, Eggeling C, Hell SW. Nat.
Methods. 2008; 5:943–945. [PubMed: 18794861]
(17). Heilemann M, van de Linde S, Mukherjee A, Sauer M. Angew. Chem., Int. Ed. 2009; 48:6903–
6908.
(18). Conley NR, Biteen JS, Moerner WE. J. Phys. Chem. B. 2008; 112:11878–11880. [PubMed:
18754575]
(19). Wombacher R, Heidbreder M, van de Linde S, Sheetz MP, Heilemann M, Cornish VW, Sauer M.
Nat. Meth. 2010; 7:717–721.
(20). Los GV, et al. ACS Chem. Biol. 2008; 3:373–382. [PubMed: 18533659]
(21). O'Hare HM, Johnsson K, Gautier A. Curr. Opin. Struct. Biol. 2007; 17:488–494. [PubMed:
17851069]
(22). So M-K, Yao H, Rao J. Biochem. Biophys. Res.Comm. 2008; 374:419–423. [PubMed:
18621022]
J Am Chem Soc. Author manuscript; available in PMC 2011 November 3.

Lee et al.
Page 5
(23). Lord SJ, Conley NR, Lee HD, Nishimura SY, Pomerantz AK, Willets KA, Lu Z, Wang H, Liu N,
Samuel R, Weber R, Semyonov A, He M, Twieg RJ, Moerner WE. ChemPhysChem. 2009;
10(1):55–65. [PubMed: 19025732]
(24). Schriven, EFV., editor. Azides and Nitrenes: Reactivity and Utility. Academic Press; Orlando,
FL: 1984.
(25). Shapiro L, McAdams HH, Losick R. Science. 2009; 326:1225–1228. [PubMed: 19965466]
(26). Biteen J, Moerner WE. Cold Spr. Harb. Persp. Biol. 2010; 2(3):a000448.
(27). Bowman G, Comolli L, Gaietta G, Fero M, Hong S-H, Jones Y, Lee J, Downing K, Ellisman M,
McAdams H, Shapiro L. Mol. Micro. 2010; 76(1):173–189.
(28). McAdams H, Shapiro L. FEBS Lett. 2009; 583:3984–3991. [PubMed: 19766635]
(29). Bernhardt TG, de Boer PA. Mol. Microbiol. 2003; 48:1171–1182. [PubMed: 12787347]
(30). Dempsey G, Bates M, Kowtoniuk WE, Liu DR, Tsien RY, Zhuang X. J. Am. Chem. Soc. 2009;
131:18192–18193. [PubMed: 19961226]
J Am Chem Soc. Author manuscript; available in PMC 2011 November 3.

Lee et al.
Page 6
Figure 1.
2
The absorption of 1 in ethanol decreases during irradiation with 1.1 mW/cm at 385 nm
shown at 15 s, 30 s, 90 s, and 150 s (left down arrow). Concurrently, the absorption of amine
photoproduct 2 grows proportionally (center up arrow). The bright fluorescence from 2
when pumped at 594 nm is the dotted curve; the heavy dashed curve is dim fluorescence
from the original sample of 1 (the small emission signal is most likely from the pre-activated
amine contaminants in the azide sample). Preactivation can be minimized by keeping the
samples in complete darkness. Compound 3 has the same photophysical properties as 1.
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Figure 2.
Evidence that the HaloTag-targeted fluorogen correctly labels specific proteins and enables
SR imaging in mammalian cells. (A) Phase image of fixed WT HeLa cells. (B) The cells in
A imaged in the DCDHFV-P channel. (C) The cells in A imaged in the Alexa488 channel.
(D) Phase image of fixed HeLa expressing HaloEnz-α-tubulin labeled with 3. (E) The cells
in D imaged in the DCDHF-V-P channel. (F) The cells in D imaged in Alexa488 channel.
(G) Overlay of E, F, and additional blue DAPI channel to show nuclei. (H) Live CHO cells
co-transfected to express both HaloEnz–α-tubulin and α-tubulin–eGFP labeled with 3 and
imaged in DCDHF-V-P channel. (I) Cells from H imaged in the EGFP channel. (The higher
background in H may be the result of nonspecific binding and imperfect washing of
untargeted fluorophores.) (J) Fixed BS-C-1 cells expressing HaloEnz-α-tubulin labeled with
3 imaged using conventional diffraction-limited imaging. Indicated microtubule measures
450±40nm FWHM. (K) Same cell as J with SR imaging. Indicated microtubule measures
85±15nm FWHM. See SI for sample preparation procedures.
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Figure 3.
Diffraction-limited imaging of 4 inside live C. crescentus cells expressing fusion proteins to
25
FtsZ and PopZ. These proteins localize as expected, indicating that the HaloTag–DCDHF
labeling does not disrupt typical cellular behavior.
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Figure 4.
SR imaging of protein fusions inside live C. crescentus cells using 3. (A–C) PopZ forms a
polymeric network at the poles of the cells. Compared to the DL images in Figure 3, these
SR images reveal distinct shapes of the PopZ structure, including the cap-like network in C.
(D–E) AmiC is recruited to the division plane early in the cell cycle. These SR images
indicate that AmiC may form a structure that hugs the membrane. For details of imaging and
image processing, see SI. The SR images are extracted from localizations over 75 seconds
with a mean localization precision of 32 ± 12 nm.
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Scheme 1.
14,15
Photochemical activation produces 2 from 1. A mixture of photoproducts is produced,
but the primary amine with R =R =H is the significant product (see Table 1 for reaction
1
2
yield of primary amine). HaloTag versions of 1 and a separate fluorophore are also shown (3
and 4).
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Table 1
Photophysical/photochemical parameters. DCDHF-V-P-azide is the earlier underivatized azido-DCDHF from
ref 9
for comparison.
[a]
[b]
[d]
Φ p
[c]
Yield
λabs,azide (nm)
λabs,amine/λfl,amine (nm)
570/613
DCDHF-V-P-azide
424
443
–
65%
~50%
–
0.0059
0.095
–
1 (and 3)
572/627
4
598/629
[a]
[b]
[c]
[d]
Peak absorbance for azido fluorogen.
Absorbance and fluorescence peak wavelengths of the amino fluorophore.
Overall chemical reaction yield to the primary fluorescent amine.
Photoconversion quantum yield of azido fluorogens to any product (i.e. the probability of photoconverting after absorbing one photon). For
−5 reference 11
reference, the value of Φ for mEos is on the order of 10 (see
for more detailed comparisons). For all DCDHFs, the fluorescence
P
quantum yield of the photoactivated form varies greatly depending on the precise nanoenvironment, typically <0.1% in buffer to >30% when
23
rigidized (e.g. in the membrane or when bound to proteins). For measurement details, see SI. Compound 3 has the same photophysical properties
as 1.
J Am Chem Soc. Author manuscript; available in PMC 2011 November 3.