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Simultaneous Quanti fi cation of Multiple Mitochondrial and Cellular Readouts

Simultaneous Quanti fi cation of Multiple Mitochondrial and Cellular Readouts

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540



360

587

504

577

504



TMRM (mitochondria)



Fura-2 (cytosol)

Indo-1 (cytosol)

Rhod-2 (mitochondria)



HEt (cytosol)/Mito-HEt

(MitoSOX) (mitochondria)

DCFDA (cytosol)



BCECF (cytosol)

SNARF®-1 (cytosol)



Autofluorescence (mitochondria)



ER-Tracker® Red (ER)

ER-Tracker® Green (ER)



LysoTracker® Red (lysosome)

LysoTracker® Green (lysosome)



Dy



Calcium (Ca2+)



ROS



pH



NADH



Endoplasmatic

reticulum



Lysosomes



592

511



615

511



480



528

640



580

535



505

405 + 485

590



590



Emission



x



x



x



x



x



x



x



mito-AcGFP1

(475/505)



x



x



x



x



x



x

x



mito-tagRFP

(555/584)



Mask generation (Em/Ex)



Invitrogen (L7528)

Invitrogen (L7526)



Invitrogen (E34250)

Invitrogen (E34251)



(31)



Invitrogen (B1150)

Invitrogen (C1272)



Invitrogen (D11347/

M36008), (32)

Invitrogen (C6827), (13)



Invitrogen (F1221), (29)

Invitrogen (I1223), (30)

Invitrogen (R1245MP), (31)



Invitrogen (T668), (17)



References



One or more probes are given for each parameter with cellular localization between parentheses and the predicted combination of the probe with either mitochondria targeted

mito-AcGFP1 or mito-tagRFP for mitochondrial mask generation. The maximum excitation and emission wavelengths of commonly used chemical fluorescent probes are given

under in vitro conditions. For optimized spectral separation, intracellular in vivo spectra should be recorded (see Sect. 5)



503

575



396 + 510

505



340 + 380

350

540



Excitation



Probe



Parameter



Wavelengths (in vitro)



Table 1

Simultaneous quantification of mitochondrial and cellular parameters using chemical fluorescent probes and mitochondria

targeted mito-AcGFP1 or mito-tagRFP fluorescent proteins

6

Live-Cell Quantification of Mitochondrial Functional Parameters

123



Ratiometric-pericam (cytosol)

Ratiometric-pericam-mt (mitochondria)

Cameleon-1 BFP-GFP (cytosol)

Cameleon-2 CFP-YFP (cytosol)



HyPer (cytosol)

MitoHyPer (mitochondria)



SypHer (cytosol)

MitoSypHer (mitochondria)



ATeam (cytosol)

Mito-ATeam (mitochondria)



Calcium (Ca2+)



ROS



pH



ATP



435



420 + 490



475 + 527



x



x

x



535/25a



x

x

x

x



mito-tagRFP

(555/584)



x

x



mito-AcGFP1

(475/505)



516



445 + 510

472 + 527



381

433

420 + 500



517



Emission



415 + 494



Excitation



Mask generation (Em/Ex)



(37)



(36)



(35)



(34)



(33)



References



For each cellular and mito- parameter, the proteinaceous sensors and its cellular localization is given between parentheses. Following the in vitro maximum excitation and emission

wavelengths, these sensors could be combined with mito-tagRFP for mitochondrial mask generation. In case of FRET-based sensors, both channels combined could function

as mitochondrial mask image. For optimized illumination, intracellular in vivo spectra should be recorded (see Sect. 5)

a

The maximum emission wavelength of SypHer is not yet known, although probably similar to HyPer. Instead, the emission filter set used in reference (36) is given



Probe



Parameter



Wavelengths (in vitro)



Table 2

Simultaneous quantification of mitochondrial and cellular parameters using proteinaceous fluorescent sensors

and mito-targeted mito-AcGFP1 or mito-tagRFP fluorescent proteins



124

M. Nooteboom et al.



6



Live-Cell Quantification of Mitochondrial Functional Parameters



125



sensor HyPer, based on the bacterial protein OxyR, and its pH

sensing synthetic mutant SypHer (35, 36). The free (ATP) can be

measured by the ATP sensor ATeam. This sensor is based on the

bacterial e subunit derived from the F0F1-ATP synthase of Bacillus

subtilis and Bacillus sp. PS3 (37). In principle, all of these reporters should be able to be combined with mito-tagRFP for

mitochondrial localization (Table 2). However, the optimal excitation and emission wavelengths given for both the chemical and

protein-based probes are generally determined under in vitro conditions. The latter might not realistically mimic the cellular environment. Therefore it is recommended to record intracellular

in vivo spectra of each fluorescent reporter to allow proper selection of excitation and emission filters for microscopy, in order to

avoid fluorescent bleed-through of either wavelengths. This can

for instance be performed using a spectrophotofluorometer (e.g.,

Shimadzu RF-5301PC, Shimadzu Scientific Instruments, Tokyo,

Japan).

Two aspects have to be considered when one intends to combine the use of two different proteinaceous fluorescent probes.

First, both probes have to be introduced into the cell. This can be

achieved either by transfection of the cells with two plasmids or by

a vector encoding both proteins. Correct protein synthesis and

protein folding has to be examined, for example using native gel

electrophoresis and in-gel fluorescence analysis (27). Second, certain ratiometric protein-based sensors are pH-sensitive. For

instance, in the case of the ATP:ADP ratio sensing reporter

Perceval, pH correction was carried out by paralleled measurements of with the pH indicator SNARF-5F (38). In case of ATeam,

its YFP/CFP emission ratio was virtually pH-independent under

physiological pH ranges (37). The individual wavelengths of ratiometric sensors can be assessed as internal control. Synchronization

of both wavelengths (i.e., increase or decrease in fluorescence signal) will indicate an environmental or morphological change,

whereas asynchronous behavior of both wavelengths is a strong

indicator of ligand sensing. Currently, so-called “dead” sensors

that are identical to the normal sensors but unable to bind the

ligand are considered the optimal controls for sensor specificity.



Acknowledgments

This work was supported by an equipment grant of NWO

(Netherlands Organization for Scientific Research, No: 911-02008), the Dutch Ministry of Economic Affairs (Innovative

Onderzoeks Projecten (IOP) Grant: #IGE05003), and by the

CSBR (Centres for Systems Biology Research) initiative from

NWO (No: CSBR09/013V).



126



M. Nooteboom et al.



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SE, Grefte S, Smeitink JA, Willems PH (2006)

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14. Condreay JP, Kost TA (2007) Baculovirus

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AS, Swarts HG, Mayatepek E, Smeitink JA,

Willems PH (2008) Life cell quantification of

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organelle level. Cytometry A 73:129–138

18. Kuznetsov AV, Hermann M, Saks V, Hengster

P, Margreiter R (2009) The cell-type specificity

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19. Koopman WJ, Distelmaier F, Esseling JJ,

Smeitink JA, Willems PH (2008) Computerassisted live cell analysis of mitochondrial membrane potential, morphology and calcium

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Localization of mitochondria in living cells with

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comparison of different modes of evaluating

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human skin fibroblasts. Cytometry A 69:1–12

27. Dieteren CE, Willems PH, Vogel RO, Swarts

HG, Fransen J, Roepman R, Crienen G, Smeitink

JA, Nijtmans LG, Koopman WJ (2008) Subunits

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Mitochondrial dynamics in human NADH:

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Biochem Cell Biol 41:1773–1782

29. Visch HJ, Rutter GA, Koopman WJ, Koenderink

JB, Verkaart S, de Groot T, Varadi A, Mitchell

KJ, van den Heuvel LP, Smeitink JA, Willems

PH (2004) Inhibition of mitochondrial

Na+-Ca2+ exchange restores agonist-induced

ATP production and Ca2+ handling in human

complex I deficiency. J Biol Chem

279:40328–40336

30. Paredes RM, Etzler JC, Watts LT, Zheng W,

Lechleiter JD (2008) Chemical calcium indicators. Methods 46:143–151

31. Visch HJ, Koopman WJ, Zeegers D, van

Emst-de Vries SE, van Kuppeveld FJ, van den

Heuvel LW, Smeitink JA, Willems PH (2006)

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32. Forkink M, Smeitink JAM, Brock R, Willems

PHGM, Koopman WJH (2010) Detection and

manipulation of mitochondrial reactive oxygen

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(2001) Circularly permuted green fluorescent

proteins engineered to sense Ca2+. Proc Natl

Acad Sci USA 98:3197–3202

34. Miyawaki A, Llopis J, Heim R, McCaffery JM,

Adams JA, Ikura M, Tsien RY (1997)

Fluorescent indicators for Ca2+ based on green

fluorescent proteins and calmodulin. Nature

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35. Belousov VV, Fradkov AF, Lukyanov KA,

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Lukyanov S (2006) Genetically encoded

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encoded fluorescent reporter of ATP:ADP

ratio. Nat Methods 6:161–166



Chapter 7

Functional Imaging Using Two-Photon Microscopy

in Living Tissue

Ivo Vanzetta, Thomas Deneux, Attila Kaszás, Gergely Katona,

and Balazs Rozsa

Abstract

Over the last 20 years, neuroscientists have become increasingly interested in two-photon microscopy. One

of the reasons for this interest is that two-photon fluorescence excitation allows counterbalancing the

deterioration of the optical signals due to light scattering, and this opens the door for high-resolution

imaging at considerable depth in living tissue. Due to progress in fluorescent marking techniques, to date,

two-photon microscopy allows the functional exploration of neuronal activity at multiple scales, from the

subprocesses of a single cell (dendrites, single spines, etc.), through single cells or small networks of a few

neurons, up to large neuronal populations in the order of a cortical column. Here, we provide some information on the practical aspects of two-photon microscopy applied to imaging neurons in living tissue.

We first discuss the advantages, shortcomings, and possible developments of the technique. We provide

some practical considerations on the choice of the microscope itself, as well as on its principal elements.

Because of recent progress in tackling high-speed imaging of 3D objects, we devote particular attention to

the discussion of z-axis scanning techniques. Next, we illustrate some common applications, such as calcium imaging of neuronal activity, in vitro and in vivo. We also briefly illustrate how two-photon microscopy can be used for the imaging of erythrocyte flow in individual capillaries. Some practical considerations

on specific protocols are provided in the form of self-consistent text boxes.

Key words: Two-photon, Laser scanning, In vivo, Fluorescence microscopy, Functional imaging,

Calcium imaging, 3D scanning, Erythrocyte movement



1. Introduction

Fluorescence microscopy has proven an essential tool for the examination of biological specimens, both fixed or alive (1). One of the

reasons for its great success is that fluorescent objects can be selectively visualized at good signal-to-background ratio even at small

fluorophore concentrations.



Emilio Badoer (ed.), Visualization Techniques: From Immunohistochemistry to Magnetic Resonance Imaging, Neuromethods,

vol. 70, DOI 10.1007/978-1-61779-897-9_7, © Springer Science+Business Media, LLC 2012



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When applied to thick (over ~100 mm) samples, conventional

fluorescent microscopy suffers under the shortcoming that the

fluorophore is excited in a comparatively large volume, giving rise

to a large out-of-focus component in the fluorescence reaching

the detector. Confocal microscopy (2, 3) deals with this problem

by using a pinhole to reject the photons originating from all locations other than from one point in the focal plane, providing an

optical section of the sample. The focal plane is then scanned in

two dimensions (x, y). The same procedure is repeated, serially, for

each desired depth (z-dimension) until the relevant part of the

biological specimen is covered. Subsequently, the imaged volume

is visualized using three-dimensional reconstruction. It is important to realize that, although in confocal microscopy the out-of-focus

photons do not contribute to the image, the corresponding molecules are still excited, leading to a high imaging cost in terms of

photodamage and photobleaching. A partial solution to this problem consists in recording through many pinholes in parallel (e.g.,

using Nipkow disk-based approaches). In addition to collecting,

for a given excitation volume, photons emitted from more than

one location, the same sample locations are visited multiple times,

leading to a higher signal rate. This allows the lowering of laser

powers used and finally leads to less photodamage and bleaching.

Another shortcoming, however, remains: detection through

the pinhole(s) blocks not only the fluorescence originating from

out-of-focus excitation, but also the photons that were indeed

emitted from the in-focus-point, yet underwent scattering on their

way out of the tissue. This loss of “good” photons further reduces

the yield of the instrument. All these problems are exacerbated by

the strong light scattering of biological tissue, which makes highresolution depth-imaging impossible with conventional and confocal microscopy.

The application of two-photon excitation to biological microscopy can thus be considered as a true breakthrough in fluorescent

microscopy depth-imaging. Several excellent reviews of two-photon

functional imaging, and calcium imaging in particular, can be

found in the literature. They either focus on its technical aspects

(4–8), on its applications (9–12), or on its place in the wider context of the various recent technological developments, which provide tools for a “reverse engineering” on the brain (13). In this

introduction, we therefore sketch only briefly the basic principles

and advantages of two-photon microscopy over more conventional

approaches.

Two-photon excitation occurs when two low-energy photons

together excite a fluorescent molecule, which then decays back to

its fundamental state by emission of a photon of somewhat lower

energy than the sum of the two exciting ones. The theoretical

grounds of two-photon absorption were identified as early as 1931

by Maria Göppert-Mayer (14). However, experimental confirmation



7



Functional Imaging Using Two-Photon Microscopy in Living Tissue



131



had to wait for progress in laser technology; in the 1960s that

progress made ultrashort pulsed lasers available, which finally

allowed the design of two-photon microscopes (15).

Why has two-photon microscopy become so popular over the

last 10 years or so?

First, living tissue scatters light less in the infrared than at visible wavelengths, at which most common fluorophores are typically

excited in single-photon mode. It thus became possible to optically

examine fluorescent objects within living specimen up to depths

that were previously inaccessible (5, 16). These reach from typical

depths of several hundreds of microns up to 1 mm (17), although

in the latter case the power of the excitation pulses has to be

boosted via regenerative amplification (18).

Second, two photons must be absorbed simultaneously for

excitation to occur. Therefore, the probability that a fluorescent

molecule is excited depends quadratically on light intensity, rather

than linearly as in the case of single-photon fluorescence. In simple

terms, significant excitation occurs only where the local concentration of photons is very high, which happens essentially only at the

focal point of the microscope, even in the presence of scattering.

Photobleaching and photodamage to the tissue are thus greatly

reduced. Most important, no fluorescence is emitted from outof-focus locations resulting in “automatic” optical sectioning,

without any need for out-of-focus rejection strategies like that

implemented in confocal microscopy by the usage of one or more

pinholes.

Finally, in two-photon microscopy all emitted fluorescence

photons are useful—even the ones that were scattered on their way

out of the tissue. Collecting them allows a dramatic increase in

signal-to-noise ratio (SNR) with respect to situations where the

scattered photons are rejected, as in confocal microscopy. Note that

multiple-scattered photons are emitted from a seemingly large field

of view (an area of ~1.5 times the recording depth, see: (19)) requiring a collection optics with a sufficiently large acceptance angle.

For the above reasons, two-photon excitation can selectively

excite (or photochemically activate or inactivate) microscopic parts

of a biological sample, at depths up to 1 mm and only at the in-focus

plane, avoiding the photodamage and photobleaching that would

result from a comparable excitation with confocal microscopy.

Overall, two-photon microscopy is thus a unique tool for

imaging biological samples—in vivo in particular—in depth, as

well as for local photochemistry. However, its performance in

imaging transparent or very thin specimen is worse than that of

confocal or even wide field fluorescence microscopy, because the

longer wavelengths used for imaging reduce the maximal achievable spatial resolution. As often in biological research, the choice of

the optimal imaging technique shall thus depend on the particular

scientific question that needs to be addressed.



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2. Materials

2.1. Microscopes:

General



(a) Build or buy?

For high-end techniques such as multiphoton fluorescence,

the instrument is obviously of fundamental importance. Twophoton microscopes have become commercially available from

most major producers, as well as from a few smaller companies,

providing the user with turn-key solutions implemented in

ready-to-go setups. “In-house” solutions sometimes constitute a valuable alternative to purchasing an all-finished instrument, both because of economical considerations and because

this allows for more flexibility in the mechanical design, which

is particularly important for in vivo applications. If it is possible

to convert confocal microscopes into two-photon ones (15), it

is somewhat cumbersome. Therefore, it is often preferable to

build a two-photon microscope essentially from scratch.

Detailed descriptions of the needed components and how to

proceed to assemble them can be found in excellent publications (e.g., (6, 7, 20, 21)). Importantly, software has also

become available that allows to reliably control laser scanning

(22, 23). Summarizing, building one’s own instrument has

the important advantage that it allows to customize the microscope to one’s specific needs. However, a good degree of experience and technical know-how should be available in-house

and/or experienced, qualified assistance is highly commendable

(but see also (24) for a user-friendly adaptation of a low-cost

commercial microscope to two-photon imaging). Figure 1

shows a typical setup for a two-photon microscope.

(b) Lasers and objectives

Excitation light must be provided by a femtosecond pulsed

laser source, which nowadays are made commercially available

by several producers. Critical parameters are wavelength, pulse

width and power, which depend to some extent on the specific

application. Available since the 1980s, Ti:sapphire lasers appear

to comply with the needs posed by most applications, warranting a more than reasonable compromise between the different

requirements. Commercial Ti:Sapphire lasers are wavelengthtunable over a large range (~690–1,040 nm), allowing to

match the excitation wavelength of a wide range of fluorescent

molecules. However, for wavelengths above 1,000 nm, as

needed for the excitation of red fluorescent proteins, Ti:sapphire

lasers perform poorly, making other light sources preferable,

such as Ti:Sapphire laser pumped optical parametric oscillators

(OPOs, ~1,000–1,600 nm), mode-locked Ytterbium-doped

lasers (25, 26) (1,030–1,060 nm, nontunable) or Cr:Forsterite

lasers (27) (tunable, 1,200–1,300 nm).



7



Functional Imaging Using Two-Photon Microscopy in Living Tissue



133



Scanning mirrors

Mode-locked

Ti:Slaser

Infrared laser

excitation



MHz repetition



Detector



dichroic

mirror



PMT



rate



~100 fs

impulse width



Visible

fluorescent

light



Coverglass

Sample



excitation localized

to the focal point



Fig. 1. A typical setup for a two-photon microscope. A MHz femtosecond infrared pulsed laser beam is sent onto a set of

galvanometric scanning mirrors, providing lateral deflection of the excitation beam, which is thus focused onto different xy

locations in the sample preparation by a high numerical aperture objective, often of the water-immersion type. Fluorescence,

in the visible domain, is then collected by the same objective and sent toward the detector, usually a PMT through a

dichroic mirror, which eliminates backscattered excitation light. The z-position (depth of focus) has to be adjusted independently, either manually, or using piezoelectric actuators (see text). Recently, it has been shown that scanning in all three

(x, y and z) dimensions can be performed without mechanical movement, using opto-acoustic deflectors (44).



Ti:Sapphire lasers are pumped with a laser beam from an

argon or frequency-doubled Nd:YVO4 laser and yield outputs

of 2 W average power or more in the central part of their wavelength range. This power is usually enough, except for ultradeep

imaging (500–1,000 mm), where the laser pulses have to be

powered-up by regenerative amplification, at the expenses of

pulse frequency (typically by a factor of 1,000 (17, 18)). Note

however, that, ultimately, a depth limit is imposed on imaging

by the fluorescence generated at the surface of the sample by

out-of-focus light, with a consequently worse localized excitation and a deterioration of contrast. In gray matter, this limit

appears to be in the order of 1,000 mm (17).

Pulse rates have to be such as to balance the fluorophore’s

excitation efficiency and onset of saturation. This criterion is

met by the typical Ti:Sapphire pulse rates (~80 MHz) for



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I. Vanzetta et al.



most common molecules, which is fortunate because in

mode-locked lasers pulse rate is difficult to change. Stable excitation rate being an important consideration, it is essential

that a constant number of pulses arrive on a pixel. Assuming a

typical pulse rate of 80 MHz and a typical dwell time of 1 ms,

more than 80 pulses arrive on a pixel. The number of pulses

per pixel will thus be stable at the 1.25% level.

The objective generates the laser focus required for localization of excitation and is therefore a critical element of the

microscope. A large choice of objectives suitable for two-photon

microscopy can be found on the market. Important criteria for

a choice include: (1) the numerical aperture, which has to be

large because it determines resolution and the solid angle

within which fluorescence is collected; (2) the magnification,

which has to be chosen according to the desired field of view;

(3) the working distance, which has to be relatively large, especially in in vivo applications where several media have to be

stacked between objective and sample (coverglass, agarose,

etc.); and (4) a good transmission efficiency in the near-IR.

Note that some of these requirements are to some extent

conflicting (for instance, increasing the working distance of an

objective without increasing its exit pupil leads to a decreased

numerical aperture), so that in most cases the optimal choice

consists in making the best compromise between the different

requirements. Given the considerable price of two-photon

microscope objectives, budgetary issues are often part of this

compromise. A good discussion of lasers and objectives can be

found in (5, 7, 11).

(c) Pulse length and power

Optimal pulse length is more difficult to define. The optical

components in the excitation path cause dispersion, pulses

reaching the sample are thus broadened (“chirped”) with

respect to their initial duration. Indeed, in an optical medium

the propagation speed of electromagnetic radiation depends

on wavelength, and this effect is stronger for short pulses (typically around or below 100 fs) than for long ones, because of

their different envelop in Fourier space. Being constant and

known for a given microscope’s excitation path, such dispersive broadening can be precompensated by “negatively dispersing” the pulses. These “prechirped” pulses are thus restored

to short ones when exiting the objective, which maximizes

two-photon absorption at the sample level (5). Although this

procedure is relatively easy to implement (28, 29), the added

optical components unavoidably result in some power loss of

which one has to take into account.

More problematic is the limitation on peak power and thus

on pulse duration posed by photobleaching and photodamage,



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Simultaneous Quanti fi cation of Multiple Mitochondrial and Cellular Readouts

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