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}
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\newline
% Insert author names, affiliations and corresponding author email (do not include titles, positions, or degrees).
\\
-Manuel Schottdorf\textsuperscript{1,*}, % 0000-0002-5468-4255
+Manuel Schottdorf\textsuperscript{1,2*}, % 0000-0002-5468-4255
P. Dylan Rich\textsuperscript{1}, % 0000-0001-9782-7984
E. Mika Diamanti\textsuperscript{1}, % 0000-0003-1199-3362
-Albert Lin\textsuperscript{1,4}, % 0000-0002-4541-5889
+Albert Lin\textsuperscript{1,3}, % 0000-0002-4541-5889
Sina Tafazoli\textsuperscript{1}, % 0000-0003-1926-0227
-Edward H. Nieh\textsuperscript{1,2}, % 0000-0003-2154-6224
-Stephan Y. Thiberge\textsuperscript{1,3*} % 0000-0002-6583-6613
+Edward H. Nieh\textsuperscript{1,4}, % 0000-0003-2154-6224
+Stephan Y. Thiberge\textsuperscript{1,5*} % 0000-0002-6583-6613
\\
\bigskip
\textbf{1} Princeton Neuroscience Institute, Princeton University, Princeton, NJ, USA\\
-\textbf{2} School of Medicine, University of Virginia, Charlottesville, VA, USA \\
-\textbf{3} Bezos Center for Neural Circuit Dynamics, Princeton University, Princeton, NJ, USA\\
-\textbf{4} Center for the Physics of Biological Function, Princeton University, Princeton, NJ, USA\\
+\textbf{2} Psychological and Brain Sciences, University of Delaware, Newark, DE, USA\\
+\textbf{3} Center for the Physics of Biological Function, Princeton University, Princeton, NJ, USA\\
+\textbf{4} Department of Pharmacology, School of Medicine, University of Virginia, Charlottesville, VA, USA \\
+\textbf{5} Bezos Center for Neural Circuit Dynamics, Princeton University, Princeton, NJ, USA\\
\bigskip
% Use the asterisk to denote corresponding authorship and provide email address in note below.
-* mschottdorf@princeton.edu\\
+* maschott@udel.edu\\
* thiberge@princeton.edu
\end{flushleft}
% Please keep the abstract below 300 words
\section*{Abstract}
-Many laboratories use two-photon microscopy through commercial suppliers, or homemade designs of considerable complexity. The integrated nature of these systems complicates customization, troubleshooting as well as grasping the principles of two-photon microscopy. Here, we present ``Twinkle'': a microscope for Two-photon Imaging in Neuroscience, and Kit for Learning and Education. It is a fully open, high-performance and cost-effective research and teaching microscope without any custom parts beyond what can be fabricated in a university machine shop. The instrument features a large field of view, using a modern objective with a long working distance and large back aperture to maximize the fluorescence signal. We document our experiences using this system as a teaching tool in several two week long workshops, exemplify scientific use cases, and conclude with a broader note on the place of our work in the growing space of open-source scientific instrumentation.
+Many laboratories use two-photon microscopy through commercial suppliers, or homemade designs of considerable complexity. The integrated nature of these systems complicates customization, troubleshooting, and \textcolor{blue}{training on the} principles of two-photon microscopy. Here, we present ``Twinkle'': a microscope for Two-photon Imaging in Neuroscience, and Kit for Learning and Education. It is a fully open, high performing and \textcolor{blue}{easy-to-set-up microscope that can effectively be used for both education and research}. The instrument features \textcolor{blue}{a $>1\text{ mm}^2$ field of view}, using a modern objective with \textcolor{blue}{3 mm} working distance and \textcolor{blue}{two inch diameter optics combined with GaAsP photomultiplier tubes} to maximize the fluorescence signal. We document our experiences using this system as a teaching tool in several two week long workshops, exemplify scientific use cases, and conclude with a broader note on the place of our work in the growing space of open-source scientific instrumentation.
\linenumbers
\section*{Introduction}
%
-Two-photon microscopy\cite{Denk1990, Svoboda1997, So2000, Helmchen2005} is a workhorse of modern systems neuroscience across species, and two-photon microscopes exist in many system neuroscience laboratories\cite{Luu2024, Grienberger2022}. For example, in the animal model \textit{Mus musculus} (mouse), two-photon microscopes have been used to classify the function and neural inventory of the retina \cite{Baden2016}, study sequential activity in cortex \cite{Harvey2012} and subcortical structures \cite{Nieh2021}, led to transformative insights into signals in the dopaminergic reward system \cite{Engelhard2019}, and revealed an organized map among grid cells \cite{Yu2018}. In non-human primates, two-photon microscopes have been used as optical brain computer interfaces \cite{Trautmann2021}, and to survey the spatial organization of motor cortex in reach movements \cite{Ebina2018}. They also have potential to further our understanding of the function of the primate retina \cite{Sharma2016, Schottdorf2021}. In Zebrafish, \textit{Danio rerio}, two-photon microscopy can image the entire brain with single cell resolution \cite{Renninger2013}. Two-photon microscopy has also become a standard method for monitoring neural activity in invertebrates. In the fruit fly, \textit{Drosophila melanogaster}, imaging from sparse, genetically specified neuron populations \cite{Seelig2010, Lin2022} has enabled many circuits to be functionally mapped, including auditory \cite{Baker2022}, courtship \cite{Deutsch2020, Roemschied2023}, and navigation circuits \cite{Kim2017}. Two-photon microscopes have also enabled volumetric pan-neuronal imaging, in which large portions of the whole fly brain can be recorded from with high temporal resolution \cite{Pacheco2021,Brezovec2024}. In \textit{Caenorhabditis elegans}, it allowed to measure an atlas of neural signal propagation \cite{Randi2023}.\newline
-However, these systems can be expensive and difficult to customize \cite{Diamanti2021}, while improvements and new inventions are published regularly, such as the incorporation of adaptive optics to improve the signal amplitude \cite{Yao2023}, the combination of two-photon imaging with two-photon stimulation for all-optical interrogation of neural circuits \cite{Nikolenko2008, Packer2012,Rickgauer2014}, and extremely large fields of view for mesoscopic imaging across brain areas \cite{Sofroniew2016}. Recently, several approaches have also been developed to increase the size of the volumes being imaged beyond the limits set by the traditional sampling strategy while keeping the time resolution constant. New approaches probing extended regions rely on signal demixing to reconstitute the underlying structure \cite{Song2017, Kazemipour2019}. Another approach uses a set of axially separated and temporally distinct foci to record the entire axial imaging range near-simultaneously \cite{Demas2021}.\newline
-The two-photon methods' popularity suggests the need for a simple platform, at reasonable cost, for practical education, dissemination, methods development and research. We developed a high-performance and cost-effective two-photon microscope that can easily be built in any neuroscience laboratory over the course of several days and can be effectively used for both teaching and research purposes. Reflecting its use for ``\textbf{TW}o-photon \textbf{I}maging in \textbf{N}euroscience, and \textbf{K}it for \textbf{L}earning and \textbf{E}ducation'', we chose the acronym Twinkle. Twinkle's open design makes the building experience ideal for teaching the principles of two-photon microscopy to the next generation of researchers. Here, we share our design, document system performance, explore possible research applications, and report our experiences collected during several teaching workshops.
+Two-photon microscopy\cite{Denk1990, Svoboda1997} \textcolor{blue}{is a key technology across the modern life sciences\cite{Zipfel2003}, for example, in physiology\cite{Ueki2020, Dunn2002}, cancer research \cite{Perry2012}, plant biology \cite{Cheung2010}, and neuroscience\cite{So2000, Helmchen2005}. New scientific use cases and technological improvements are published at a remarkable rate\cite{Luu2024}.} However, these instruments \textcolor{blue}{are often published without detailed building instructions and explanations}. This suggests the need for an \textcolor{blue}{easy-to-set-up microscope, at reasonable cost, that can be effectively used for education, dissemination, methods development and research.} We developed a high-performance and cost-effective two-photon microscope that can \textcolor{blue}{easily be produced} in many neuroscience laboratories. Reflecting its use for ``\textbf{TW}o-photon \textbf{I}maging in \textbf{N}euroscience, and \textbf{K}it for \textbf{L}earning and \textbf{E}ducation'', we chose the acronym Twinkle. In this article, we share our design, document the performance of the system, explore possible research applications, and report our experiences collected during several teaching workshops. \textcolor{blue}{Complete CAD drawings, bill of materials, optics simulations, and detailed building instructions are provided in the supplement.}
\section*{Materials and methods}
-In two-photon laser-scanning microscopy\cite{Denk1990}, laser light is focused to a small excitation volume, the ``focal point'', which is moved across the sample by the intermediary of resonant and galvanometric scanning mirrors. As the focal point probes different locations, different quantities of fluorescence signal are detected. An image is then formed, mapping the detector photocurrent values to the locations of excitation volume: low and high photocurrents becoming respectively dark and bright pixels. Central to two-photon fluorescence excitation is the light source: a femtosecond-pulsed infrared laser of sufficient pulse energy to achieve, when focused by an objective lens, a photon density high enough for simultaneous absorption of two photons by a fluorescent molecule \cite{Denk1990, Svoboda1997, So2000, Helmchen2005}. Here, we design an open and high-performing microscope that uses these principles. As such we have made all optical and mechanical designs, electronics, bill of materials, CAD assembly, testing results, and relevant schematics open to the public. Excluding laser and optical table, the cost is around US-\$ 110k (2024). A narrative of the design and high-level view is presented here. For further technical details, and complete built documentation, we refer the reader to \nameref{S1_Appendix}, and the repository at \url{https://github.com/BrainCOGS/Microscope}.
+In two-photon laser-scanning microscopy\cite{Denk1990}, laser light is focused to a small excitation volume, the ``focal point'', which is moved across the sample by the intermediary of resonant and galvanometric scanning mirrors. As the focal point probes different locations, different quantities of fluorescence signal are detected. An image is then formed, mapping the detector photocurrent values to the locations of excitation volume: low and high photocurrents, respectively, becoming dark and bright pixels. Central to two-photon fluorescence excitation is the light source: a femtosecond-pulsed infrared laser of sufficient pulse energy to achieve, when focused by an objective lens, a photon density high enough for simultaneous absorption of two photons by a fluorescent molecule. Here, we design an open and high-performing microscope that uses these principles. As such, we have made all optical and mechanical designs, electronics, bill of materials, CAD assembly, testing results, and relevant schematics open to the public. \textcolor{blue}{Excluding laser and optical table, the cost is around US-\$ 110k (2024). Including laser and table, the cost is around US-\$ 190k (2024).} A narrative of the design and the high-level view is presented here. For further details, we refer the reader to \nameref{S1_Appendix}. All code, figures and text, CAD designs, optics simulations, and BOM are available at \url{https://github.com/BrainCOGS/Microscope}.
\subsection*{Design specifications}
To summarize our design goals, we aimed for a mechanical and optical assembly using as many off-the-shelf components as possible, with only few custom aluminium parts that can be machined in any university machine shop. Our design facilitates the adaptation for \invivo imaging by providing a large space for the organism and ancillary hardware. Our system is made cost-effective, in part, by the availability of femtosecond laser systems based on fiber technology. Operating at a fixed wavelength, these systems come at the fraction of the cost of a tunable Ti:Sapphire laser \cite{Bueno2019}.\newline
We aimed for (1) $\gtrapprox20\text{ cm}$ of free space around the objective in all directions to aid integration of the microscope with various peripheral equipment such as behavior recording and stimulation, and the sample stage, and (2) imaging on a $\approx 700\times 700\text{ \textmu m}^2$ minimum size field of view which is typical for imaging brain tissue at cellular resolution. (3) Compared to earliest open designs\cite{Rosenegger2014, Mayrhofer2015}, our system can make use of the large Numerical Aperture (NA) of modern long-working-distance objectives to collect more fluorescence, but requires to adapt our design to their larger back apertures \cite{Janelia2024}.\newline
-An overview of the microscope is shown in Fig.~\ref{fig1}. Fig.~\ref{fig1}A shows the system as a cartoon. Full details will be provided below. In short, we chose a Spark Alcor 920 nm femtosecond pulsed laser, whose beam diameter is extended by a $2\times$ magnification telescope, before it reaches the scanning mirrors. The scanning mirrors are purchased as a mount-assembled set. Located as close as possible to each other, as we will see in the alignment procedure, neither of them can be exactly positioned in the focal plane of the scan lens. The choice of mounted scanners simplifies the design and construction, but it also leads to some optical distortions across the field of view. We found that this distortion does not exceed a few percent in our design (see below). The combined scan and tube lenses magnify the beam further to fill the back aperture of the objective. Fluorescence from the specimen is then collected and brought to the aperture of two GaAsP photomultiplier tubes with suitable color filters. Fig.~\ref{fig1}B shows the CAD design of the entire assembly with important parts of the optical path labelled. Fig.~\ref{fig1}C shows an annotated picture of the microscope head. Notice that compared to the cartoon in Fig.~\ref{fig1}A, the optical path is folded with several mirrors. These mirrors make the microscope assembly more compact.
+An overview of the microscope is shown in Fig.~\ref{fig1}. Fig.~\ref{fig1}A,B shows the system as a cartoon. Full details will be provided below. In short, we chose a Spark Alcor 920 nm femtosecond pulsed laser, whose beam diameter is extended by a $2\times$ magnification telescope, before it reaches the scanning mirrors. The scanning mirrors are purchased as a mount-assembled set. Located as close as possible to each other, as we will see in the alignment procedure, neither of them can be exactly positioned in the focal plane of the scan lens. The choice of mounted scanners simplifies the design and construction, but it also leads to some optical distortions across the field of view. \textcolor{blue}{We found that this distortion does not exceed 5\% in our design (see below)}. The combined scan and tube lenses magnify the beam further to fill the back aperture of the objective. Fluorescence from the specimen is then collected and brought to the aperture of two GaAsP photomultiplier tubes with suitable color filters. Fig.~\ref{fig1}C shows the CAD design of the entire assembly with important parts of the optical path labeled. Fig.~\ref{fig1}D shows an annotated picture of the microscope head. Notice that compared to the cartoon in Fig.~\ref{fig1}A,B, the optical path is folded with several mirrors. These mirrors make the microscope assembly more compact.
\begin{figure}[!t]
\includegraphics[width=\textwidth]{fig1.jpg}
- \caption{{\bf System overview.} \textbf{A)} Cartoon of the layout. \textbf{B)} CAD drawing of the system on an optical table with several key components in the optical path highlighted. The spacing between two holes on the table is 1 inch. \textbf{C)} Photograph of the microscope head with key elements labelled. This section is visible in panel B.}
+ \caption{{\bf System overview.} \textbf{A)} \textcolor{blue}{Key components of the microscope head. Shaded colors illustrate beam deflection angles. \textbf{B)} Cartoon of the beam conditioning components. First the beam is split by 50\%, the intensity adjusted with an electro-optic modulator (EOM) and the beam width with a telescope before entering the microscope head. \textbf{C)} 3D drawing of the full microscope, highlighting beam conditioning subsystem (blue) and the microscope head (yellow). For scale, the spacing between two holes on the optical table is 1 inch or 2.5 cm. \textbf{D)} Photograph of the microscope head with key elements labeled. This section is visible in panel A.}}
\label{fig1}
\end{figure}
\subsection*{Assembly on the table}
-Various components are used between the laser and the microscope head to condition the beam, and for intensity control, Fig.~\ref{fig1}B. Out of the laser head, the beam first encounters a waveplate (AHWP10M-980, Thorlabs). This is a birefringent crystal that can rotate the linear polarization of the laser to any angle. Next, the beam travels through a polarizing beamsplitter cube (PBS103, Thorlabs). This splits the beam into two orthogonal polarization components whose relative intensity is adjusted by the orientation of the waveplate. We set the orientation of the waveplate to split the beam into equal power components, the second one being eventually used for a second microscope installed on the other extremity of the table. In the CAD drawing, this second beam is sent to a beam block (LB1, Thorlabs). Next, the beam passes through an electro-optic modulator, a Pockels cell (350-80-LA-02, Conoptics). This device allows rapid and electronic control of the laser's power level. It uses a crystal whose refractive index is controlled by an external electric field, combined with a polarizer. This can be thought of as a voltage-controlled wave plate, in which the electric field controls how much light travels through the polarizer. Next, the beam travels through an open mechanical 6 mm diameter shutter (LS6S2ZM1, Vincent Associates). When closed, the laser reflected on the shutter reflective blades is sent to another beam block. Beam blocks are not perfect absorber, so we can use advantageously the weak back scattered light from the beam block to calibrate the Pockels cell. In our set-up, a photodiode (PDA36A2, Thorlabs) is thus facing the beam block. Following the shutter, the beam travels though a $2\times$ telescope (GBE02-B, Thorlabs) doubling its diameter. Past the telescope, the beam is then reflected off several mirrors in a periscope configuration before entering a custom aluminium box housing, mounted to a 95 mm optical rail (XT95 series, Thorlabs), that houses the two scanning mirrors (CRS8K/6215H scanning mirror set, Novanta), shown in blue in Fig.~\ref{fig1}C. Leaving the scanners, the beam then travels first through the scan lens (green) and tube lens (red), passes through the long pass dichroic mirror of the collection box assembly (yellow) and finally the objective (cyan). The design of these components is covered in the next paragraphs. The steering mirrors in our system are all P01 silver protected mirrors from Thorlabs. Silver mirrors contribute minimally to dispersion which is inevitable when light travels through various optical components. Limiting the dispersion makes it possible to correct for it with the laser's built-in dispersion compensation.
+Various components are used between the laser and the microscope head to condition the beam, and for intensity control, Fig.~\ref{fig1}B. Out of the laser head, the beam first encounters a waveplate (AHWP10M-980, Thorlabs). This is a birefringent crystal that can rotate the linear polarization of the laser to any angle. Next, the beam travels through a polarizing beamsplitter cube (PBS103, Thorlabs). This splits the beam into two orthogonal polarization components whose relative intensity is adjusted by the orientation of the waveplate. We set the orientation of the waveplate to split the beam into equal power components, the second one being eventually used for a second microscope installed on the other extremity of the table. In the cartoon, this second beam is sent to a beam block (LB1, Thorlabs). \textcolor{blue}{We want to stress that this beam-splitting hardware is optional. However, modern lasers have enough power to supply two microscopes at the same time. It is therefore often strategic to split the beam into two beams, with half of the intensity each, to allow one laser (the most expensive component) to power two systems.} Next, the beam passes through an electro-optic modulator (EOM), a Pockels cell (350-80-LA-02, Conoptics). This device allows rapid and electronic control of the laser's power level. It uses a crystal whose refractive index is controlled by an external electric field, combined with a polarizer. This can be thought of as a voltage-controlled wave plate, in which the electric field controls how much light travels through the polarizer. Next, the beam travels through an open mechanical 6 mm diameter shutter (LS6S2ZM1, Vincent Associates). When closed, the laser reflected on the shutter reflective blades is sent to another beam block. Beam blocks are not perfect absorber, so we can use advantageously the weak back scattered light from the beam block to calibrate the Pockels cell. In our set-up, a photodiode (PDA36A2, Thorlabs) is thus facing the beam block. Following the shutter, the beam travels though a $2\times$ telescope (GBE02-B, Thorlabs) doubling its diameter. Past the telescope, the beam is then reflected off several mirrors in a periscope configuration before entering a custom aluminium box housing, mounted to a 95 mm optical rail (XT95 series, Thorlabs), that houses the two scanning mirrors (CRS8K/6215H scanning mirror set, Novanta), shown in blue in Fig.~\ref{fig1}D. Leaving the scanners, the beam then travels first through the scan lens (green) and tube lens (red), passes through the long pass dichroic mirror of the collection box assembly (yellow) and finally the objective (cyan). The design of these components is covered in the next paragraphs. The steering mirrors in our system are all P01 silver protected mirrors from Thorlabs. Silver mirrors contribute minimally to dispersion which is inevitable when light travels through various optical components. Limiting the dispersion makes it possible to correct for it with the laser's built-in dispersion compensation.
\subsection*{Optical design: Scan and tube lens}
-Like for all modern microscopes, infinity-corrected objectives are being used in this design. The incident laser light has to be collimated at the back aperture of the objective. For this reason, scan and tube lens must be carefully configured as a telescope. Additionally, to guaranty the laser beam enters the objective back aperture with the same amount of clipping, the scanning mirrors must be positioned in the conjugated plane of objective back aperture. In many commercial infinity corrected microscope designs, the distance between the tube lens and the objective can be chosen within a certain range. In our design, we added the constraint of positioning the objective and tube lens in a telescope, i.e. the distance between them, $d_{TO}$, is the sum of the tube lens and objective's focal length, $d_{TO} = f_O + f_T$. While this has no real impact, it prepares the assembly for possible upgrade where methods such as remote focusing could be used \cite{Botcherby2007, Botcherby2012, Sofroniew2016}.
-Our design was initiated by choosing a proper focal length for the custom assembly serving as a scan lens. We settled on a focal length $f_S=100\text{ mm}$ scan lens built from two inch diameter components \cite{Yao2023}. This choice was influenced by the availability of off-the-shelf lenses and the specific scanning range of the scanners (typically $\pm10\text{ deg}$, cf. Fig.~\ref{fig1}A). To fill the back aperture of the objective with our laser beam, we aimed for a combined magnification of $M=3.75\times=f_T/f_S$, making the focal length of the tube lens $f_T=375\text{ mm}$. With these choices, the resulting scanning angles at the back aperture of the objective are $\pm10\text{ deg}/3.75\approx\pm2.7\text{ deg}$ and with our chosen objective (Nikon $16\times$ with $f_O=12.5\text{ mm}$) the expected approximate field of view diameter is $\approx 1.2\text{ mm}$).\newline
+Like for all modern microscopes, infinity-corrected objectives are being used in this design, see Fig.~\ref{fig1}A. The incident laser light has to be collimated at the back aperture of the objective. For this reason, scan and tube lens must be carefully configured as a telescope. Additionally, to guaranty the laser beam enters the objective back aperture with the same amount of clipping, the scanning mirrors must be positioned in the conjugated plane of objective back aperture. In many commercial infinity corrected microscope designs, the distance between the tube lens and the objective can be chosen within a certain range. In our design, we added the constraint of positioning the objective and tube lens in a telescope, i.e. the distance between them, $d_{TO}$, is the sum of the tube lens and objective's focal length, $d_{TO} = f_O + f_T$. While this has no real impact, it prepares the assembly for possible upgrade where methods such as remote focusing could be used \cite{Botcherby2007, Botcherby2012, Sofroniew2016}.
+Our design was initiated by choosing a proper focal length for the custom assembly serving as a scan lens. We settled on a focal length $f_S=100\text{ mm}$ scan lens built from two inch diameter components \cite{Yao2023}. This choice was influenced by the availability of off-the-shelf lenses and the specific scanning range of the scanners (typically $\pm10\text{ deg}$, cf. Fig.~\ref{fig1}A). To fill the back aperture of the objective with our laser beam, we aimed for a combined magnification of $M=3.75\times=f_T/f_S$, making the focal length of the tube lens $f_T=375\text{ mm}$. With these choices, the resulting scanning angles at the back aperture of the objective are $\pm10\text{ deg}/3.75\approx\pm2.7\text{ deg}$ and with our chosen objective (Nikon $16\times$ with $f_O=12.5\text{ mm}$) the expected approximate field of view diameter is $\approx 1.2\text{ mm}$).\newline
The tube lens was also assembled as a lens group from off-the-shelf parts. The relative positions of the components in the two lens groups were optimized in Ansys Zemax OpticStudio. Details are provided below.
%
\begin{figure}[!t]
\includegraphics[width=\textwidth]{fig2.jpg}
- \caption{{\bf Optical design of the excitation pathway.} \textbf{A)} Zemax design with the scanning mirror, tube and scan lens groups. Colors indicate light rays produced by different mirror angles of the scanning mirror. Notice how the rays converge in the image plane. \textbf{B)} CAD design built around the optical design in A, see also Fig.~\ref{fig1}B. \textbf{C)} Cuts through the scan and tube lens assemblies, which are fabricated from off-the-shelf lenses housed in SM2 lens tubes. The specific lenses are stated in the text. \textbf{D)} Simulation of the optical system's two-photon point spread function (PSF-2p) for 0 deg deflection angle, and two example radial sections. \textbf{E)} PSF-2p in the xz plane across the full range of deflection angles. Shown below is Petzval field curvature (blue), and radial (purple) and axial (yellow) full-width-at-half-maximum of the PSF-2p. Error bars are resolution limits of the Huygens PSF estimates in sequential mode.}
+ \caption{{\bf Optical design of the excitation pathway.} \textbf{A)} Zemax design with the scanning mirror, tube and scan lens groups. Colors indicate light rays produced by different mirror angles of the scanning mirror. Notice how the rays converge in the image plane. \textbf{B)} CAD design built around the optical design in A, see also Fig.~\ref{fig1}C. \textbf{C)} Cuts through the scan and tube lens assemblies, which are fabricated from off-the-shelf lenses housed in SM2 lens tubes. The specific lenses are stated in the text. \textbf{D)} Simulation of the optical system's two-photon point spread function (PSF-2p) for 0 deg deflection angle, and two example radial sections. \textbf{E)} PSF-2p in the xz plane across the full range of deflection angles. Shown below is Petzval field curvature (blue), and radial (purple) and axial (yellow) full-width-at-half-maximum of the PSF-2p. Error bars are resolution limits of the Huygens PSF estimates in sequential mode.}
\label{fig2}
\end{figure}
%
-Fig.~\ref{fig2}A shows the propagation of light rays for various deflection angles of the scanning mirrors, and how the beams converge on a line under the objective. Note that in our simulation, in the absence of a model for our commercial objective, we replaced it by a perfect geometrical lens. Once the separation between optical elements was optimized, a model was exported in CAD (Autodesk Inventor and Fusion 360), and the optomechanics was designed around it, see Fig.~\ref{fig2}B. The CAD design and lens groups are highlighted in Fig.~\ref{fig2}B,C (see Fig.~\ref{fig1}C of a photograph of the same part of the instrument). The key properties that we optimized when designing the excitation system was an essentially flat, and diffraction limited focal point across the deflection angle range of the scanning mirrors. Fig.~\ref{fig2}D shows an estimate of the two-photon point spread function (PSF-2p), computed as the square of the Huygens PSF in Zemax. Analyzing the PSF-2p across optical deflection angles, Fig.~\ref{fig2}E, suggests very small field curvature if the objective used behaves close to a perfect geometrical lens. In this simulation, the axial resolution was $\delta z = 5.0\text{ \textmu m}$, and the radial resolution $\delta r = 0.75\text{ \textmu m}$ across a wide range of deflection angles.\newline
+Fig.~\ref{fig2}A shows the propagation of light rays for various deflection angles of the scanning mirrors, and how the beams converge on a line under the objective. Note that in our simulation, in the absence of a model for our commercial objective, we replaced it by a perfect geometrical lens. Once the separation between optical elements was optimized, a model was exported in CAD (Autodesk Inventor and Fusion 360), and the optomechanics was designed around it, see Fig.~\ref{fig2}B. The CAD design and lens groups are highlighted in Fig.~\ref{fig2}B,C (see Fig.~\ref{fig1}D of a photograph of the same part of the instrument). The key properties that we optimized when designing the excitation system was an essentially flat, and diffraction limited focal point across the deflection angle range of the scanning mirrors. Fig.~\ref{fig2}D shows an estimate of the two-photon point spread function (PSF-2p), computed as the square of the Huygens PSF in Zemax. Analyzing the PSF-2p across optical deflection angles, Fig.~\ref{fig2}E, suggests very small field curvature if the objective used behaves close to a perfect geometrical lens. In this simulation, the axial resolution was $\delta z = 5.0\text{ \textmu m}$, and the radial resolution $\delta r = 0.75\text{ \textmu m}$ across a wide range of deflection angles.\newline
The specific elements that allowed for this performance were as follows: For the scan lens, we used an assembly of four lenses, see Fig.~\ref{fig2}C, 1: KPC070AR (Newport); 2: LB1199-B (Thorlabs) and 3\&4: $2\times$ ACT508-200-B (Thorlabs). The $2\times$ ACT508-200-B lenses are arranged back-to-back and provide the majority of the optical power. The symmetric Pl\"ossl design corrects for the odd aberrations: coma, distortion, and lateral colors \cite{Negrean2014, Kidger2001}. The two additional lenses provide corrections to the even aberrations: spherical aberration, astigmatism, and field curvature. We chose the specific lenses and their spacing based on availability from established optics suppliers and the simulation of dozens of combinations in Zemax. The tube lens is inspired by a Petzval lens design \cite{Smith2007, Kidger2001}, consisting of a pair of achromatic doublets with the same orientation, $2\times$ ACT508-750-A, Thorlabs, a design performing better than a Pl\"ossl pair as observed by others as well \cite{Hong2022, Bumstead2018, Mayrhofer2015}.\newline
The two lens groups combined provide a magnification of $M=3.75\times$ to produce a beam that slightly underfills the back aperture of the Nikon NA 0.8 $16\times$ LWD objective, effectively reducing its excitation NA to $\approx0.7$. This can be advantageous for \invivo Calcium imaging in certain conditions (see discussion). Of further note, we used a piezoelectrical collar to mount the objective. The collar can move the objective along the optical axis at a fast rate for imaging across multiple planes.
@@ -153,9 +152,9 @@ \subsection*{Optical design: Collection optics}
We aimed to record fluorescence from two well-separated spectral ranges, a green ($525\pm25\text{ nm}$) and a red channel ($600\pm25\text{ nm}$). Zemax was used to select the lenses of the collection unit and their relative position, while attempting to maximize the collection of light by the detectors. A cartoon is shown in Fig.~\ref{fig3}A. The colors illustrate different point sources of fluorescent light in the sample plane. Notice the mild divergence, and approximately collimated beams that enter the aperture of the photomultiplier tube. As previously, this design was exported into CAD, and the optomechanics assembled around it, see Fig.~\ref{fig3}B. Fig.~\ref{fig3}C shows a simulation in Zemax demonstrating that our design brings nearly all the light escaping the objective back aperture with an angle below $\approx8\text{ deg}$ to the aperture of the photomultiplier tubes.
\subsection*{Mechanical design}
-Our microscope was mounted on a 16 inch thick optical table. The design was done in CAD Inventor and Fusion 360 (Autodesk), and the results shown in Fig.~\ref{fig1}B. The aim was a mechanical assembly that uses as few custom parts as possible. This is relatively easy to achieve for the excitation path, but hardly practical for the collection system which require to be both efficient and light tight. The components shown in the CAD files can be assembled and aligned by an experienced researcher in a few days. When used for teaching, a careful assembly and alignment is viable with about a week of work. Regarding the footprint, the system can comfortably be built on an $4\times4$ ft area of table. The microscope documented here was setup on a $4\times8$ ft table, shared with a second microscope. The beam-splitting hardware is shown in Fig.~\ref{fig1}B as well.\newline
+Our microscope was mounted on a 16 inch thick optical table. The design was done in CAD Inventor and Fusion 360 (Autodesk), and the results shown in Fig.~\ref{fig1}C. The aim was a mechanical assembly that uses as few custom parts as possible. This is relatively easy to achieve for the excitation path, but hardly practical for the collection system which require to be both efficient and light tight. The components shown in the CAD files can be assembled and aligned by an experienced researcher in a few days. When used for teaching, a careful assembly and alignment is viable with about a week of work. \textcolor{blue}{When set up on an imperial-unit table, the system can comfortably be built on $4\times4 \text{ ft}^2$. The microscope documented here was built on a $4\times8\text{ ft}^2$ ft table, shared with a second microscope. In metric units, our microscope has a footprint of $\approx 1.5\text{ m}^2$. In other words, a standard $1.2\times2.5\text{ m}^2$ table can house two of the systems described here.} The beam-splitting hardware \textcolor{blue}{used for this arrangement} is shown in Fig.~\ref{fig1}B,C as well.\newline
The custom mechanical components (e.g. the light-tight aluminium housing of the collection optics) were built in the university's machine shop from aluminium stock or Thorlabs parts. Some parts have tight tolerances which suggests manufacturing in a professional machine shop is preferred. If useful for training, this could however be done by students. An example are the aluminium parts that hold the large dichroic mirrors and optical filters in the collection optics. After drilling the apertures for light to pass through, little metal is left on the part which can potentially lead to warping and distortions if not machined with precision.\newline
-Finally, we want to emphasize the difference between a theoretically optimal design in Zemax and finite tolerances in real-world optomechanical parts. It is key for a good microscope design to be robust against such errors. In our design, the objective is fixed, while the mirrors, the tube and scan lens assemblies, and the scanning mirrors can be re-positioned along the optical axis for alignment. This allows the mechanical components sufficient degrees of freedom to optically align all key parts of the microscope. In the supplement, \nameref{S1_Appendix}, we go through this alignment procedure in detail.
+Finally, we want to emphasize the difference between a theoretically optimal design in Zemax and finite tolerances in real-world optomechanical parts. It is key for a good microscope design to be robust against such errors. In our design, the objective is fixed, while the mirrors, the tube and scan lens assemblies, and the scanning mirrors can be re-positioned along the optical axis for alignment. This allows the mechanical components sufficient degrees of freedom to optically align all key parts of the microscope. In the supplement, we go through this alignment procedure in detail.
\subsection*{Scanners power supplies and controllers enclosure}
The scanning mirrors are active components. They are controlled by two circuit boards supplied with external power. The driver board of the resonant scanner does not dissipate much heat, but the driver of the slow galvanometer can become warm in normal operation. This heat needs to be dissipated. We mount the circuit boards in a custom electronics box (Gold Box, Acopian), making sure to use heat transfer compound between the circuit boards and the aluminium box, and to mount them securely together with screws. At time of purchase, we requested the controller of the slow galvanometer to be configured for usage at $\pm 28\text{ V}$ (the manufacturer default is $\pm 24\text{ V}$ ). This overall improves dynamical properties. The controller requires three voltage inputs, respectively -28 V, 0 V, and +28 V. The 0 V is also grounded. The second power supply provides 12 V for the resonant scanner. It is beneficial to choose a power supply that can stably produce these voltages for years. We have made good experiences with Agilent E3630A, and the Instek GPD-3303D. Importantly, double check the power supplies with a good multimeter before connecting the driver boards. Please also see \nameref{S1_Appendix} for a detailed illustration.
@@ -171,17 +170,15 @@ \subsection*{PMTs control circuit}
A simple circuit, see Fig.~\ref{fig4}A, is used to control and display the photomultiplier gain. The H16201P-40 photomultiplier tube module comes with four cables, two for power supply, and two for gain control. These leads are color-coded. A small circuit for resistance programming of the gain was designed which also displays the gain setting voltage on a small digital panel meter (Murata DMS-20PC). The circuit for controlling two channels (red and green) is shown in Fig.~\ref{fig4}A. For the tube module, the +15 V supply voltage and ground are provided via the black and red cables. The blue cable is connected to ground via a potentiometer to act as a voltage divider. The gain voltage is then fed back to the module via the white input line, and used for display on the panel meter. A 12k resistor between the PMT module and the potentiometer was added to limit the control voltage which is safer for the PMT. Optimal signal-to-noise ratio for the PMTs is usually $\approx 0.7-0.8\text{ V}$. The panel meter needs an additional +5 V supply that we obtain from the +15 V via a resistive divider. The meter is operating in single-ended input configuration, see Fig.~\ref{fig4}B. Due to the simplicity of this circuit, we do not solder this on a circuit board, but rather in a prototype development box directly, see Fig.~\ref{fig4}C, which is attached to the rig. This circuit needs a well regulated power supply. For convenience, we also use an Agilent E3630A. The PMT outputs were amplified with two Transimpedance Amplifiers (TIA60, Thorlabs), and directly connected to the data acquisition system.
\subsection*{Ancillary hardware}
-Our microscope is controlled with ScanImage \cite{Pologruto2003} running on a Windows PC equipped with a data acquisition system (DAQ), both from the same supplier (MBF Bioscience).\newline
-Typical Ti:Sapphire tunable femtosecond pulsed laser source can be expensive. For the work demonstrated here, we used a fixed-wavelength femtosecond laser systems, based on fiber technology (e.g.\cite{Bueno2019,Limpert2006,Wise2012,Young2015}). These systems come at a fraction of the cost, and they take up significantly less real estate on an optical table. The laser was controlled with the same PC as ScanImage through a USB connection. As the beam travels through a number of crystals, lenses, and mirrors with various chromatic properties, significant dispersion is introduced which increases pulse duration, reducing the two-photon absorption efficiency. It is possible to compensate for the group-delay dispersion of the microscope by giving the short-wavelength components a sufficient head start so that blue and red components arrive at the sample at the same time. We found, empirically, that the built-in group velocity dispersion compensation of the laser was more than sufficient to tune the system to an optimum, which we found close to $\varphi=-20200\pm170\text{ fs}^2$. Details of this measurement, and some theory, are provided below as a teaching example (cf. Fig.~\ref{fig9}). Regarding light transmission, we found the Spark laser to produce $\approx 2.1\text{ W}$ of light. After the half-wave plate and beamsplitter cube, $\approx 1.0\text{ W}$ enter the Pockels cell (cf. Fig.~\ref{fig1}B). The Pockels cell is rotated so that the control voltage produces the largest dynamic range of transmitted laser power (we go through the alignment procedure in depth below). At maximum transmission setting of the Pockels cell, we obtain $\approx 260\text{ mW}$ below the objective, a large value, as it is exceeding the maximum recommended power for cortical imaging in mice \cite{Song2017}. All pictures presented in this article were obtained with $\approx 15\text{ mW}$ below the objective.
+Our microscope is controlled with ScanImage \cite{Pologruto2003} running on a Windows PC equipped with a data acquisition system (DAQ), both from the same supplier (MBF Bioscience). For the work demonstrated here, we used a fixed-wavelength femtosecond laser systems, based on fiber technology (e.g.\cite{Bueno2019,Limpert2006,Wise2012,Young2015}). \textcolor{blue}{These devices come at $\approx 1/3$ of the cost of a Ti:Sapphire system, and take up significantly less real estate on the optical table.} Our laser, \textcolor{blue}{an Alcor 920 manufactured by Spark Lasers}, was controlled with the same PC as ScanImage through a USB connection. As the beam travels through a number of crystals, lenses, and mirrors with various chromatic properties, significant dispersion is introduced which increases pulse duration, reducing the two-photon absorption efficiency. It is possible to compensate for the group-delay dispersion of the microscope by giving the short-wavelength components a sufficient head start so that blue and red components arrive at the sample at the same time. We found, empirically, that the built-in group velocity dispersion compensation of the laser was more than sufficient to tune the system to an optimum, which we found close to $\varphi=-20200\pm170\text{ fs}^2$. Details of this measurement, and some theory, are provided below as a teaching example (cf. Fig.~\ref{fig9}). Regarding light transmission, we found the Spark laser to produce $\approx 2.1\text{ W}$ of light. After the half-wave plate and beamsplitter cube, $\approx 1.0\text{ W}$ enter the Pockels cell (cf. Fig.~\ref{fig1}B). The Pockels cell is rotated so that the control voltage produces the largest dynamic range of transmitted laser power (we go through the alignment procedure in depth below). At maximum transmission setting of the Pockels cell, we obtain $\approx 260\text{ mW}$ below the objective, a large value, as it is exceeding the maximum recommended power for cortical imaging in mice \cite{Song2017}. All pictures presented in this article were obtained with $\approx 15\text{ mW}$ below the objective.
-\subsection*{Animal procedure}
-All procedures performed in this study were approved by the Institutional Animal Care and Use Committee at Princeton University and were performed in accordance with the Guide for the Care and Use of Laboratory Animals \cite{Guide2011}.
+\subsection*{Ethics statement}
+\textcolor{blue}{Some of the data presented here were obtained from animals. Our work was conducted according to internationally accepted standards and we followed all laws and regulations about animal research in the United States and Germany. We used vertebrate animals in the form of GCaMP6f expressing transgenic Zebrafish (\textit{Danio rerio}) for \invivo imaging. Fish were bred and housed at the Princeton Neuroscience Institute. We used five day old larvae immobilized in 2\% low melting point Agarose in E3 medium. Larvae were sacrificed after the imaging sessions. As required by law, we have obtained prior approval by the Institutional Animal Care and Use Committee at Princeton University under Protocol Number 2033. Further \invivo imaging work was done in the adult fruit fly (\textit{Drosophila melanogaster}). In the United States, work in lower-level invertebrate species like the fruit fly does not require approval by the Institutional Animal Care and Use Committee. We also imaged a previously prepared fixed and stained histological sample obtained from a Wistar rat (\textit{Rattus norvegicus}). This specific sample was made by one of us during their PhD\cite{Schottdorf2018}. The animal was bred in the animal house of the Max Planck Institute for Experimental Medicine according to European and German guidelines for experimental animals, and work was carried out with authorization of the responsible federal state authority. Finally, we imaged plant material from the common Dandilion (\textit{Taraxacum officinale}) \textit{in vivo}. This experiment does not require ethics approval in the United States.}
\section*{Results}
\subsection*{System performance}
%
-Here, we document the performance of the microscope and a few example applications using a Nikon NA 0.8 16$\times$ LWD objective (N16XLWD-PF, Thorlabs). We first determined the size and properties of the field of view with a 100~\textmu m grid (R1L3S3P, Thorlabs), imaged using a Fluorescein in water film through a standard \#1.5 ($\approx170\text{ \textmu m}$ thick) coverslip, see Fig.~\ref{fig5}A. The field of view exceeded $\approx 1\text{ mm}^2$ ($d\approx1.3\text{ mm}$ along the diagonal), and the field curvature was below the thickness of the thin Fluorescein film $\lesssim 10\text{ \textmu m}$. The maximum scan angle was limited by vignetting of the two inch diameter optics (we discuss this further in the teaching section). When measuring the local scale in \textmu m/px at low magnification across the field of view, we observed small deviations of $\Delta S \approx 0.1\text{ \textmu m/px}$ in the center when compared to the edges, while the overall range of magnifications across the field of view was tight, see Fig.~\ref{fig5}B. The average scale is $S=2.77\pm0.04\text{ \textmu m/px}$, which suggests a deviation from uniformity of $\Delta S/S = 0.1/2.8 \approx 4\%$.
%
\begin{figure}[t]
\includegraphics[width=\textwidth]{fig5.jpg}
@@ -189,7 +186,7 @@ \subsection*{System performance}
\label{fig5}
\end{figure}
%
-Next, we imaged a uniform bath of Fluorescein, see Fig.~\ref{fig5}C. This measures the quality of excitation and the efficiency of the collection optics across the field of view. Within the central $700\text{ \textmu m} \times 700\text{ \textmu m}$ region, the signal deviated from uniformity within $\approx 13\%$ ($\pm 1 \text{ standard deviation}$), see Fig.~\ref{fig5}D, E. Following these measurements, we imaged a bead sample of 0.2\microns diameter in 1\% Agarose (Dragon Green beads; Bangs Laboratories) to measure axial and radial PSFs. The sample is shown in Fig.~\ref{fig5}F. Averaging across the $N=38$ beads in this volume, we measured a radially symmetric full-width-at-half-maximum of $\delta r\approx760\pm30\text{ nm}$, see Fig.~\ref{fig5}G,H, and axially $\delta z \approx 5.4\pm0.9\text{ \textmu m}$, see Fig.~\ref{fig5}I. The theoretical resolution limits\cite{Tsai2002} for our system, underfilled to $\text{NA}=0.7$ at $\lambda=920\text{ nm}$ and water immersion ($n=1.33$) were radially FWHM of $\delta r=0.6\,\lambda/\text{NA}=780\text{ nm}$ and axially $\delta z = 2\,\lambda n/\text{NA}^2=5.0\text{ \textmu m}$. This is consistent with our earlier simulations in Zemax (cf. Fig.~\ref{fig2}), and suggests that our microscope is operating very close to the diffraction limit, and its design specifications.
+Here, we document the performance of the microscope and a few example applications using a Nikon NA 0.8 16$\times$ LWD objective (N16XLWD-PF, Thorlabs). We first determined the size and properties of the field of view with a 100~\textmu m grid (R1L3S3P, Thorlabs), imaged using a Fluorescein in water film through a standard \#1.5 ($\approx170\text{ \textmu m}$ thick) coverslip, see Fig.~\ref{fig5}A. The field of view exceeded $\approx 1\text{ mm}^2$ ($d\approx1.3\text{ mm}$ along the diagonal), and the field curvature was below the thickness of the thin Fluorescein film $\lesssim 10\text{ \textmu m}$. The maximum scan angle was limited by vignetting of the two inch diameter optics (we discuss this further in the teaching section). When measuring the local scale in \textmu m/px at low magnification across the field of view, we observed small deviations of $\Delta S \approx 0.1\text{ \textmu m/px}$ in the center when compared to the edges, while the overall range of magnifications across the field of view was tight, see Fig.~\ref{fig5}B. The average scale is $S=2.77\pm0.04\text{ \textmu m/px}$, which suggests a deviation from uniformity of $\Delta S/S = 0.1/2.8 \approx 4\%$. Next, we imaged a uniform bath of Fluorescein, see Fig.~\ref{fig5}C. This measures the quality of excitation and the efficiency of the collection optics across the field of view. Within the central $700\text{ \textmu m} \times 700\text{ \textmu m}$ region, the signal deviated from uniformity within $\approx 13\%$ ($\pm 1 \text{ standard deviation}$), see Fig.~\ref{fig5}D, E. Following these measurements, we imaged a bead sample of 0.2\microns diameter in 1\% Agarose (Dragon Green beads; Bangs Laboratories) to measure axial and radial PSFs. The sample is shown in Fig.~\ref{fig5}F. Averaging across the $N=38$ beads in this volume, we measured a radially symmetric full-width-at-half-maximum of $\delta r\approx760\pm30\text{ nm}$, see Fig.~\ref{fig5}G,H, and axially $\delta z \approx 5.4\pm0.9\text{ \textmu m}$, see Fig.~\ref{fig5}I. The theoretical resolution limits\cite{Tsai2002} for our system, underfilled to $\text{NA}=0.7$ at $\lambda=920\text{ nm}$ and water immersion ($n=1.33$) were radially FWHM of $\delta r=0.6\,\lambda/\text{NA}=780\text{ nm}$ and axially $\delta z = 2\,\lambda n/\text{NA}^2=5.0\text{ \textmu m}$. This is consistent with our earlier simulations in Zemax (cf. Fig.~\ref{fig2}), and suggests that our microscope is operating very close to the diffraction limit, and its design specifications.
\subsection*{Imaging performance}
@@ -199,12 +196,12 @@ \subsection*{Imaging performance}
\label{fig6}
\end{figure}
-Confident that the system operates as intended, we next imaged a number of representative samples that are readily available to us, namely plant material, a histological and a strongly scattering sample, see Fig.~\ref{fig6}. The first samples was a Dandilion flower (Taraxacum officinale) \cite{Rupprecht2018}. Autofluorescence in plant material is a good test to separate compounds fluorescing in the red and green channels respectively, while also exhibiting complex three-dimensional structure \cite{Cheung2010, Nguyen2001}. Fig.~\ref{fig6}A shows a slice through part of the flower, that reveals red-fluorescent structural plant material, together with green-fluorescent Dandilion pollen grains. The complex 3-D structure is readily apparent if one zooms into a small region (white box), and records a stack, see Fig.~\ref{fig6}B. The pollen grains are hidden under a layer of red-fluorescent plant material as well, which we can easily resolve. Pollen grains are a great sample in a teaching lab, as they are easy to obtain, easy to prepare, and complex when imaged with a two-photon microscope\cite{Rupprecht2018}. The next sample is a mounted primary rat cell culture\cite{Schottdorf2018}, stained for GFAP (Rabbit-Anti-GFAP, ab33922) in the green channel (Donkey-Anti-Rabbit Alexa Fluor 488, ab150061), see Fig.~\ref{fig6}C. This demonstrates the complex structure of glial fibrillary acidic protein in astrocytes. Finally, we imaged a piece of a cellulose napkin (Kimwipe, Fisher Scientific), see Fig.~\ref{fig6}D. This is an example of a strongly scattering sample with complex 3D fibrous structure. To demonstrate this structure, we acquired a volume and color-coded depth. Individual slices at different depths reveal uncorrelated fiber structures. In contrast to our earlier experiments in a less scattering sample (cf. Fig.~\ref{fig5}), the image quality here deteriorates with depth.\newline
+We next imaged a number of representative samples that are readily available to us, namely plant material, a histological and a strongly scattering sample, see Fig.~\ref{fig6}. The first samples was a Dandilion flower (Taraxacum officinale) \cite{Rupprecht2018}. Autofluorescence in plant material is a good test to separate compounds fluorescing in the red and green channels respectively, while also exhibiting complex three-dimensional structure \cite{Cheung2010, Nguyen2001}. Fig.~\ref{fig6}A shows a slice through part of the flower, that reveals red-fluorescent structural plant material, together with green-fluorescent Dandilion pollen grains. The complex 3-D structure is readily apparent if one zooms into a small region (white box), and records a stack, see Fig.~\ref{fig6}B. The pollen grains are hidden under a layer of red-fluorescent plant material as well, which we can easily resolve. Pollen grains are a great sample in a teaching lab, as they are easy to obtain, easy to prepare, and complex when imaged with a two-photon microscope\cite{Rupprecht2018}. The next sample is a mounted primary rat cell culture\cite{Schottdorf2018}, stained for GFAP (Rabbit-Anti-GFAP, ab33922) in the green channel (Donkey-Anti-Rabbit Alexa Fluor 488, ab150061), see Fig.~\ref{fig6}C. This demonstrates the complex structure of glial fibrillary acidic protein in astrocytes. Finally, we imaged a piece of a cellulose napkin (Kimwipe, Fisher Scientific), see Fig.~\ref{fig6}D. This is an example of a strongly scattering sample with complex 3D fibrous structure. To demonstrate this structure, we acquired a volume and color-coded depth. Individual slices at different depths reveal uncorrelated fiber structures. In contrast to our earlier experiments in a less scattering sample (cf. Fig.~\ref{fig5}), the image quality here deteriorates with depth.\newline
Next, we imaged neural activity \textit{in vivo} in both larval zebrafish (Danio rerio) and adult \textit{Drosophila melanogaster}.
%
\begin{figure}[t]
\includegraphics[width=\textwidth]{fig7.jpg}
- \caption{{\bf Example Calcium imaging in transgenic Zebrafish (Danio rerio).} \textbf{A)} 3D rendering of a 300~\textmu m deep volumetric image of a 5 day old Zebrafish larva with Fiji\cite{Schindelin2019}. \textbf{B)} Dorsal slice through the same animal and two zoom views down to cellular scale. \textbf{C)} Color-coded $\approx10\text{ min}$ of time-averaged volumetric imaging data of three 20~\textmu m-spaced planes using the piezoelectric collar attached to the objective. The layers are colored in red/green/blue. No motion correction was applied. Notice the blurry averages, caused by motion of the fish. \textbf{D)} Running Suite2p on the data in C produces numerous regions of interest (ROIs) with complex Calcium dynamics. Left: Suite2p applies motion correction, resulting in a much crisper average picture. Right: Time series of a few ROIs over 10 min.}
+ \caption{{\bf Example Calcium imaging in transgenic Zebrafish (Danio rerio).} \textbf{A)} 3D rendering of a 300~\textmu m deep volumetric image of a five day old Zebrafish larva with Fiji\cite{Schindelin2019}. \textbf{B)} Dorsal slice through the same animal and two zoom views down to cellular scale. \textbf{C)} Color-coded $\approx10\text{ min}$ of time-averaged volumetric imaging data of three 20~\textmu m-spaced planes using the piezoelectric collar attached to the objective. The layers are colored in red/green/blue. No motion correction was applied. Notice the blurry averages, caused by motion of the fish. \textbf{D)} Running Suite2p on the data in C produces numerous regions of interest (ROIs) with complex Calcium dynamics. Left: Suite2p applies motion correction, resulting in a much crisper average picture. Right: Time series of a few ROIs over 10 min.}
\label{fig7}
\end{figure}
%
@@ -222,8 +219,8 @@ \subsection*{Imaging performance}
\end{figure}
\subsection*{Teaching}
-Twinkle was assembled, disassembled, and reassembled three times by different groups of researchers and students. In the following paragraphs, we summarize our experiences, and produce two examples for what we consider useful learning experiences for two-photon microscopy.\newline
-To organize our workshop, we freed around 2 weeks in the summer. On the first day, we provided an introductory lecture going through the principles of two-photon microscopy (cf. Fig.~\ref{fig1}A), safety, and the plan ahead. Next, we completely stripped the optical table and returned Twinkle's components back into their containers. Then reassembly began, first setting up and aligning laser and beam-splitting hardware, then the Pockels cell, the shutter, telescope, and associated beam blocks (cf. Fig.~\ref{fig1}B). This was done over the course of days 1-3 during which students became familiar working with safety goggles, and aligning lasers using a power meter and cards. In particular, the alignment of the Pockels cell requires care, as its orientation has to be carefully set to produce the largest dynamic range of transmitted laser power depending on the control voltage (see teaching examples below). Following the conditioning of the beam, we then proceeded to assemble the mechanics of the microscope head (cf. Fig.~\ref{fig1}C). Reassembling the box housing the scanning mirrors, the lens groups from individual elements, and the collection optics was done from days 3-6 (cf. Fig.~\ref{fig2}, \ref{fig3}, \ref{fig4}) After this was done, we assessed the performance of the instrument and fine tuned it on days 7-8 (cf. Fig.~\ref{fig5}). When properly aligned, we shielded the optical path with sheet metal and Thorlabs tubing, and imaged various samples on day 9 (cf. Figs.~\ref{fig6}, \ref{fig7}, \ref{fig8}). On the last day, day 10, we addressed questions that came up during the course of the workshop.\newline
+Twinkle was assembled, disassembled, and reassembled three times by different groups of researchers and students. In the following paragraphs, we summarize our experiences, and produce two examples for what we consider useful learning experiences for two-photon microscopy. \textcolor{blue}{The detailed building instructions and additional information are available the supplement, \nameref{S1_Appendix}}.\newline
+To organize our workshop, we freed around 2 weeks in the summer. On the first day, we provided an introductory lecture going through the principles of two-photon microscopy (cf. Fig.~\ref{fig1}A), safety, and the plan ahead. Next, we completely stripped the optical table and returned Twinkle's components back into their containers. Then reassembly began, first setting up and aligning laser and beam-splitting hardware, then the Pockels cell, the shutter, telescope, and associated beam blocks (cf. Fig.~\ref{fig1}B,C). This was done over the course of days 1-3 during which students became familiar working with safety goggles, and aligning lasers using a power meter and cards. In particular, the alignment of the Pockels cell requires care, as its orientation has to be carefully set to produce the largest dynamic range of transmitted laser power depending on the control voltage (see teaching examples below). Following the conditioning of the beam, we then proceeded to assemble the mechanics of the microscope head (cf. Fig.~\ref{fig1}D). Reassembling the box housing the scanning mirrors, the lens groups from individual elements, and the collection optics was done from days 3-6 (cf. Fig.~\ref{fig2}, \ref{fig3}, \ref{fig4}) After this was done, we assessed the performance of the instrument and fine tuned it on days 7-8 (cf. Fig.~\ref{fig5}). When properly aligned, we shielded the optical path with sheet metal and Thorlabs tubing, and imaged various samples on day 9 (cf. Figs.~\ref{fig6}, \ref{fig7}, \ref{fig8}). On the last day, day 10, we addressed questions that came up during the course of the workshop.\newline
To exemplify a simple teaching example, see Fig.~\ref{fig9}. Fig.~\ref{fig9}A shows light transmission through the Pockels cell for two different orientations (red and blue) to teach the working principles of the Pockels effect. The optimal angle is shown by the blue dots. A fit to the expected transmitted power curve,
\begin{eqnarray}
P(V) = P_0 \sin\left(2\pi\frac{V}{V_0} + \phi \right)^2
@@ -240,60 +237,69 @@ \subsection*{Teaching}
\begin{eqnarray}
I(\varphi) =I_0 \left(1+\alpha\frac{(\varphi+\varphi_0)^2}{\Delta\tau^4}\right)^{-1/2}.
\end{eqnarray}
-Here, $\alpha=16\log(2)^2$. This equation can be derived by expressing the Gaussian pulse shape in the Fourier domain, applying a phase-shift along the laser path to quadratic order in the frequency, and computing the result in the time domain. Deriving this expression is a fun at-home exercise. The fit suggests that the various optical components introduced a dispersion of $\varphi_0\approx20200\pm170\text{ fs}^2$. When removed with the built-in dispersion compensation of the laser, our system operates with a FWHM pulse of $\Delta\tau \approx 150\pm20\text{ fs}$, which is close to, but significantly longer than the $\approx110\text{ fs}$ pulses produced by the laser itself (measured with an autocorrelator). This suggests detectable non-linear dispersion that we cannot compensate with our optics. Note that analyses like these, relating laser physics and fluorescence, are close to the research frontier\cite{Saidi2023}.
+Here, $\alpha=16\log(2)^2$. This equation can be derived by expressing the Gaussian pulse shape in the Fourier domain, applying a phase-shift along the laser path to quadratic order in the frequency, and computing the result in the time domain. Deriving this expression is a fun at-home exercise. The fit suggests that the various optical components introduced a dispersion of $\varphi_0\approx20200\pm170\text{ fs}^2$. When removed with the built-in dispersion compensation of the laser, our system operates with a FWHM pulse of $\Delta\tau \approx 150\pm20\text{ fs}$, which is close to, but significantly longer than the $\approx110\text{ fs}$ pulses produced by the laser itself (measured with an autocorrelator). This suggests detectable non-linear dispersion that we cannot compensate with our optics. Note that analyses like these, relating laser physics and fluorescence, are close to the research frontier\cite{Saidi2023}.
\section*{Discussion}
-In this article, we have covered the design, building and alignment of a two-photon microscope. We then assessed its performance, and demonstrated several possible use cases, from plant physiology over material science to neuroscience. In the following lines, we will expand on issues raised in passing earlier, and conclude with a note about our instrument in the growing space of open source scientific instrumentation.
+In this article, we have covered the design, building, and alignment of a two-photon microscope. We then assessed its performance and demonstrated several possible use cases, from plant physiology over material science to neuroscience. \textcolor{blue}{In the following paragraphs, we will expand on some recent scientific use cases of two-photon microscopy, discuss improvements to the design, elaborate on its use for education,} and conclude with a note about our instrument in the growing space of open source scientific instrumentation.
-\subsection*{Improvements to the microscope}
+\subsection*{Scientific use cases}
+\textcolor{blue}{Based on its unique ability to image with high resolution and limited photo damage deep into scattering samples, two-photon microscopy became an important tool across the life sciences. For example, \textit{ex vivo}, two-photon microscopy can monitor, in three dimensions, both the living and cornified keratinocytes of the epidermis, the collagen/elastin fibers in the dermal layer of the skin \cite{So1998} or the dynamics of quiescent skeletal muscle stem cells embedded in the muscle at cellular resolution \cite{Ma2024}. \textit{In vivo}, two-photon microscopy allows to monitor the dynamics of viral infections in the lung \cite{Ueki2020}, cell vitality and apoptosis, fluid transport, receptor-mediated endocytosis, blood flow, and leukocyte trafficking in the kidney \cite{Dunn2002},
+blood vascular hemodynamics in the thymus \cite{Seyedhassantehrani2024}, stem cells in the skin \cite{Huang2017}, produces new insights into cancer \textit{ex vivo} and \textit{in vivo} \cite{Perry2012}, and even entered clinical practice \cite{Konig2007}.\newline
+Arguably, two-photon microscope is particularly transformative in systems neuroscience because it facilitates the non-invasive measurement of fast neuronal dynamics with single cell resolution in awake and behaving animals \cite{Luu2024, Grienberger2022}. In the mouse, \textit{Mus musculus}, two-photon microscopes have been used to classify the function and neural inventory of the retina \cite{Baden2016}, study sequential activity in cortex \cite{Harvey2012} and subcortical structures \cite{Nieh2021}, led to transformative insights into signals in the dopaminergic reward system \cite{Engelhard2019}, and revealed an organized map among grid cells \cite{Yu2018}. In non-human primates, two-photon microscopes have been used as optical brain computer interfaces \cite{Trautmann2021}, and to survey the spatial organization of motor cortex in reach movements \cite{Ebina2018}. They also have potential to further our understanding of the function of the primate retina \cite{Sharma2016, Schottdorf2021}. In the Zebrafish larva, \textit{Danio rerio}, two-photon microscopy can image the entire brain with single cell resolution \cite{Renninger2013}. Two-photon microscopy has also become a standard method for monitoring neural activity in invertebrates. In the fruit fly, \textit{Drosophila melanogaster}, imaging from sparse, genetically specified neuron populations \cite{Seelig2010, Lin2022} has enabled many circuits to be functionally mapped, including auditory \cite{Baker2022}, courtship \cite{Deutsch2020, Roemschied2023}, and navigation circuits \cite{Kim2017}. Two-photon microscopes have also enabled volumetric pan-neuronal imaging, in which large portions of the whole fly brain can be recorded from with high temporal resolution \cite{Pacheco2021,Brezovec2024}. In \textit{Caenorhabditis elegans}, it allowed to measure an atlas of neural signal propagation \cite{Randi2023}.\newline
+New inventions are published regularly, such as the incorporation of adaptive optics to improve the signal amplitude \cite{Yao2023}, the combination of two-photon imaging with two-photon stimulation for all-optical interrogation of neural circuits \cite{Nikolenko2008, Packer2012,Rickgauer2014}, and extremely large fields of view for mesoscopic imaging across brain areas \cite{Sofroniew2016}. Recently, several approaches have also been developed to increase the size of the volumes being imaged beyond the limits set by the traditional sampling strategy while keeping the time resolution constant. New approaches probing extended regions rely on signal demixing to reconstitute the underlying structure \cite{Song2017, Kazemipour2019}. Another approach uses a set of axially separated and temporally distinct foci to record the entire axial imaging range near-simultaneously \cite{Demas2021}. We hope that our work here can contribute further momentum to these exciting new research avenues.}
-(1) We used silver mirrors to direct the beam. These reflect around 98\% of incoming light. Dielectric mirrors perform significantly better, typically exceeding 99.5\% reflectivity. With around 8 to 10 mirrors in the beam, this can boost overall transmission by 10\%-20\%, but comes at slightly higher cost and can introduce complex dispersion \cite{Rupprecht2019}. In our experiments, we have not encountered power limitations: For the data presented here, we used only $\approx 15\text{ mW}$ under the objective while at maximum transmission the system allows for up to $\approx 260\text{ mW}$ to leave the objective. For simplicity, and similar to other designs \cite{Sofroniew2016}, we therefore used exclusively silver mirrors. However, for stimulation or ablation experiments, considerations might be different. \newline
-(2) In our design, a piezoelectric collar is attached to the objective. Physically moving the objective along the optical axis allows to image several planes. The time required by the piezoelectric collar to stabilize can reach the scale of 10 ms when using large and heavy objectives. In these cases, volumetric imaging is best achieved using a ramp where the objective is constantly moving at the same speed, rather than doing steps of constant height for every planes. An electrically tunable lens (ETL) can be an alternative to the piezoobjective. These lenses have a variable focal length, usually from around 10 cm to infinity, that can be set with a control voltage within $<1\text{ ms}$. \newline
+\subsection*{Improvements/modifications to the microscope}
+(1) We used silver mirrors to direct the beam. These reflect around 98\% of incoming light. Dielectric mirrors perform significantly better, typically exceeding 99.5\% reflectivity. With around 8 to 10 mirrors in the beam, this can boost overall transmission by 10\%-20\%, but comes at slightly higher cost and can introduce complex dispersion \cite{Rupprecht2019}. In our experiments, we have not encountered power limitations: For the data presented here, we used only $\approx 15\text{ mW}$ under the objective while at maximum transmission the system allows for up to $\approx 260\text{ mW}$ to leave the objective. For simplicity, and similar to other designs \cite{Sofroniew2016}, we therefore used exclusively silver mirrors. However, for stimulation or ablation experiments, considerations might be different.\newline
+(2) In our design, a piezoelectric collar is attached to the objective. Physically moving the objective along the optical axis allows to image several planes for fast \textcolor{blue}{volumetric imaging \invivo or tissue samples \textit{ex vivo}}. The time required by the piezoelectric collar to stabilize can reach the scale of 10 ms when using large and heavy objectives. In these cases, volumetric imaging is best achieved using a ramp where the objective is constantly moving at the same speed, rather than doing steps of constant height for every planes. An electrically tunable lens (ETL) \cite{Chen2016} or \textcolor{blue}{remote focusing} \cite{Botcherby2007} are an alternative to the piezoelectric collar. \textcolor{blue}{These techniques are necessary when the objective is difficult to move (such as a very heavy objective) or cannot be moved, for example during volumetric imaging combined with controlled stimulation, as moving the objective would also change the location of the stimulation spots.}\newline
(3) Estimating the pulse duration suggests non-linear dispersion that we cannot correct. This is highly dependent on the laser. For readers interested in this aspect of microscope design, we refer to \cite{Saidi2023, Bueno2019}. If available, other light sources might be worth exploring.\newline
-(4) We made the design choice to use a resonant and slow scanning mirror pair positioned close together in the same mount. Rather than the mirrors themselves, it is the plane equidistant between the rotational axis of both scan mirrors that is conjugated to the back aperture of the objective. This is a compromise that is used by many (e.g. Thorlabs' Cerna or earlier designs \cite{Rosenegger2014, Tan1999, Nguyen2001}). Alternative microscope designs add a pair of lenses between the slow and resonant scanning mirrors to position them in conjugated planes. Our design choice significantly simplifies the design and reduces the amount of glass between laser and sample to reduce dispersion. It does, however, introduce a slight Barrel distortion that becomes visible at low magnifications.\newline
+(4) We made the design choice to use a resonant and slow scanning mirror pair positioned close together in the same mount. Rather than the mirrors themselves, it is the plane equidistant between the rotational axis of both scan mirrors that is conjugated to the back aperture of the objective. This is a compromise that is used by many (e.g. Thorlabs' Cerna or earlier designs \cite{Rosenegger2014, Tan1999, Nguyen2001}). Alternative microscope designs add a pair of lenses between the slow and resonant scanning mirrors to position them in conjugated planes. Our design choice significantly simplifies the design and reduces the amount of glass between laser and sample to reduce dispersion. It does, however, introduce a slight Barrel distortion that becomes visible at low magnifications.\newline
(5) The fixed $2\times$ beam expander can be replaced with variable/zoom beam expander that allows a user to dial in a particular beam magnification to tweak the point spread function for a particular application\cite{Smith2019}. While a high numerical aperture is beneficial for high axial resolution and improved separation of somatic and neuropil signals \cite{Kerr2008, Gobel2007}, the large angles at which excitation light is brought into the brain results in longer paths for the light to take. This can lead to more scattering. For experiments aiming to image deep in the brain, a lower NA can be beneficial \cite{Tung2004}. Similarly, the presence of axial brain motion of several \textmu m in awake and behaving animals can introduce movement artifacts at high numerical apertures \cite{Dombeck2007}. That said, even when underfilled by the excitation beam, a large NA of the objective is beneficial to collect as much fluorescent light as possible.\newline
-(6) It can be interesting to some to combine the two-photon system with wide-field imaging. To this end, the two inch mirror above the collection optics can be replaced with a 800 or 900 nm short-pass filter followed by a tube lens and a camera. This would allow infrared imaging around 700 nm through the collection optics, as it is transparent above 665 nm. While this is relatively straightforward to do \cite{Rosenegger2014}, widefield epipfluorescence is harder to realize.\newline
-(7) Our fluorescence filters have an average optical density $>4$ in the range between 400 nm and 460 nm. This is advantageous for use in virtual reality systems, because light leaving a projector can be filtered to this range before projected on a dome surrounding the objective and animal. We have successfully used budget friendly colored glass filters in front of the projector (Schott BG25 or Schott BG5) to block projector light from entering the excitation path \cite{Dombeck2007}. Combined with a light shield around the objective \cite{Dombeck2010}, such a projection system will only minimally interfere with imaging (e.g. \cite{Nieh2021}). This is a useful range for visual experiments in mice, but if other model organisms are desired, the optical filters need to be adjusted accordingly.\newline
-(8) Parts of our microscope can be replaced with commercial products. For example, we have used commercial telecentric scan lenses in the past \cite{Song2017, Rich2024} in combination with a three inch diameter tube lens. These come at higher cost and introduce higher dispersion, but exhibit very low field curvature on a very large field of view. While low field curvature is not always considered the highest priority in neuroscience applications, this can be a useful substitution.\newline
-(9) To aid mechanical stability, the aluminium plate forming the base of the head of the microscope could be replaced with a steel plate, or a thicker piece of aluminium, that is less prone to vibrations when bumped into. This would also be important if one wanted to extend the microscope head to provide even more space around the objective.
+\textcolor{blue}{(6) When imaging fixed samples, movement artifacts are not an issue. For such experiments, particularly \textit{ex vivo} work, it would be strategic to fully fill the back aperture of the objective and produce a PSF that is as tight as possible.}\newline
+\textcolor{blue}{(7) Using an objective with an even larger numerical aperture can significantly increase collection efficiency. Our objective choice was made with long working distance in mind. Our $16\times$ water-immersion objective with a numerical aperture of 0.8 accepts around $36\%$ of the $4\pi$ square radiants of emission. A possible alternative, the Nikon $25\times$ with a slightly smaller 2 mm working distance, offers a numerical aperture of 1.10 and would collect $68\%$ of light; close to a $2\times$ improvement to collection efficiency.}\newline
+\textcolor{blue}{(8) Our microscope exhibits (small) field curvature. In neuroscience, field curvature is often not considered critical because samples are rarely flat. If the microscope was used for precision length measurements, for example in \textit{ex vivo} tissue samples or for lithography purposes, one would want to optimize field flatness. Very small field curvature can be obtained with commercial f-theta scan lenses.}\newline
+\textcolor{blue}{(9) Resonant scanning mirrors are fast, but cannot be steered. If a user was interested in imaging custom ROIs, the scanning mirror pair could be replaced with a pair of slow galvanometic mirrors, which can be controlled with high precision.}\newline
+(10) For some applications, it can be beneficial to combine the two-photon system with wide-field imaging. To this end, the two inch mirror above the collection optics can be replaced with a 800 or 900 nm short-pass filter followed by a tube lens and a camera. This would allow infrared (IR) imaging around 700 nm through the collection optics, as it is transparent above 665 nm. \textcolor{blue}{Infrared illumination can be provided with an LED and a semitransparent mirror. This mirror could conveniently be housed in a flip mount to be removed from the optical path when the system is used for 2p imaging.} While this is relatively straightforward to implement \cite{Rosenegger2014}, wide-field fluorescence microscopy is harder to realize.\newline
+(11) Our fluorescence filters have an average optical density $>4$ in the range between 400 nm and 460 nm. This is advantageous for use in virtual reality systems, because light leaving a projector can be filtered to this range before projected on a dome surrounding the objective and animal. We have successfully used budget friendly colored glass filters in front of the projector (Schott BG25 or Schott BG5) to block projector light from entering the excitation path \cite{Dombeck2007}. Combined with a light shield around the objective \cite{Dombeck2010}, such a projection system will only minimally interfere with imaging (e.g. \cite{Nieh2021}). This is a useful range for visual experiments in mice, but if other model organisms are desired, the optical filters need to be adjusted accordingly.\newline
+(12) Parts of our microscope can be replaced with commercial products. For example, we have used commercial telecentric scan lenses in the past \cite{Song2017, Rich2024} in combination with a three inch diameter tube lens. These come at higher cost and introduce higher dispersion, but exhibit very low field curvature on a very large field of view. While low field curvature is not always considered the highest priority in neuroscience applications, this can be a useful substitution.\newline
+(13) To aid mechanical stability, the aluminium plate forming the base of the head of the microscope could be replaced with a steel plate, or a thicker piece of aluminium, that is less prone to vibrations when bumped into. This would also be important if one wanted to extend the microscope head to provide even more space around the objective.
\subsection*{Teaching advanced methods in neuroscience}
In addition to training researchers on how to set up a two-photon microscope, we aimed to provide examples of imaging data that are not neuroscience related, and do not require the handling of animals or living neurons (cf. Fig.~\ref{fig5}, \ref{fig6}, \ref{fig9}). Other samples are possible as well (such as cheese \cite{Nguyen2001}). Working with such samples can make experiments in a teaching lab easier as such spaces are not always designated animal work areas. If students can work with animals, the instrument is ideally suited for various neuroscience experiments. For example, the Calcium imaging demonstrated in Figs.~\ref{fig7}, \ref{fig8} could be combined with pharmacology to demonstrate changes of neural activity patterns. Or students can learn in a controlled environment how to get optical access to the nervous system and hone in their surgery skills to produce good quality optical windows\cite{Holtmaat2009, Grienberger2022}.
Our microscope was set up in a teaching lab space at the Princeton Neuroscience Institute. It was disassembled and re-assembled by various generations of students and postdocs. We found that assembling and aligning the microscope, as described, takes around 10 work-days. These two weeks seem to be a good compromise between depth, and researcher commitment. Various additional content across difficulty levels can be added depending on student interest and abilities (cf. Fig.~\ref{fig9}). For example, and only mentioned in passing before, the history and current applications of the two-photon effect and its use in microscopy is a fascinating subject to explore, e.g. \cite{Sheppard2020, Yao2023}.\newline
-Students might also become interested in further optimizing the optical design, or wonder why many lenses are used in lens groups. Questions like these can sometimes be linked to prior student knowledge. For example, for students interested in astronomy, the tube lens can be thought of as an eye piece with a long focal length and eye relief \cite{Smith2007}. In our system, the tube lens projects the image formed by the scan lens to an exit pupil, the back aperture of the objective. The objective then focuses the collimated beams onto the sample. In astronomy, the eye piece forms collimated beams of the image formed by the objective lens. The beams converge at some distance (the ``eye relief'') to enter the pupil of the eye of a human observer. The lens in the human eye then focuses the collimated beams onto the retina. Pl\"ossl designs in astronomy are known to perform particularly well for long eye relief \cite{Smith2007, Nagler1983}, and are a common design in two-photon microscopes as well (e.g. \cite{Rupprecht2024}). Our design is close to this, but flipped one of the two lenses to a Petzval configuration. In our hands, this configuration performed better \cite{Hong2022, Bumstead2018, Mayrhofer2015}.\newline
-For students interested in photography, the scan lens can be thought of as a 100 mm f/2 camera lens. Camera lenses face similar challenges as our scan lens. They have to focus distant objects, at different angles, into the same flat photographic emulsion, while faithfully depicting true angular distances with minimal aberrations. Think for example about stars on the sky, whose light arrives as beams at different angles at the camera lens. Addressing this problem lead to lens designs that were very similar to our scan lens. One important design, sometimes referred to as a Gauss doublet objective \cite{Kidger2001}, features two symmetrically configured doublets providing much of the optical power, while additional lenses control aberrations. For example, the Zeiss Planar lens from 1897 features 6 elements in 4 groups with a symmetric pair of doublets in a design very similar to our scan lens assembly \cite{Rudolf1897}. To help students and researchers explore these sometime challenging subjects theoretically, we provide all Zemax files for the microscope optics. More advanced optimization in Zemax requires the use of their programming API. For example, the simulated point spread functions and their analysis in Fig.~\ref{fig2} were computed in Matlab, using the Zemax API. The code is provided in our repository. This allows for fast optimization of system parameters by effectively scanning through lens spacing, lens types, and the like. This can allow researchers and students to explore the limits and possible future improvements of the system. Experimental modifications of the system are simpler in the format of a workshop. For example, students can swap or rotate lenses and observe the effects. The scan lens assembly is particularly sensitive to this, because the incoming laser beam is concentrated into a small region that gets scanned across the surface of the lens. Small surface imperfections will introduce optical difference across spot positions, and significantly affect field flatness, cf. Fig.~\ref{fig5}C,D,E.\newline
+Students might also become interested in further optimizing the optical design, or wonder why many lenses are \textcolor{blue}{doublets and} used in lens groups. \textcolor{blue}{Some of these questions have intuitive answers. Doublets, for example, contain more glass surfaces that can be optimized to produce a better performing lens when compared to the two surfaces of a singlet\cite{ThorlabsLens}. To our knowledge, the first mentioning of the minimization of aberration by placement of several subsequent doublets was done by Joseph Lister in 1830 \cite{Lister1830}. This idea was used for many designs, including pairs of doublets in Pl\"ossl and Petzval eye pieces, or objectives themselves \cite{Zhang2019}}. Other questions can sometimes be linked to prior student knowledge. For example, for students interested in astronomy, the tube lens can be thought of as an eye piece with a long focal length and eye relief \cite{Smith2007}. In our system, the tube lens projects the image formed by the scan lens to an exit pupil, the back aperture of the objective. The objective then focuses the collimated beams onto the sample. In astronomy, the eye piece forms collimated beams of the image formed by the objective lens. The beams converge at some distance (the ``eye relief'') to enter the pupil of the eye of a human observer. The lens in the human eye then focuses the collimated beams onto the retina. The aforementioned Pl\"ossl designs are known to perform particularly well for long eye relief in astronomy \cite{Smith2007, Nagler1983}, and are a common design in two-photon microscopes as well (e.g. \cite{Rupprecht2024}). Our design is close to this, but flipped one of the two lenses to a Petzval configuration. In our hands, this configuration performed better \cite{Hong2022, Bumstead2018, Mayrhofer2015}.\newline
+For students interested in photography, the scan lens can be thought of as a 100 mm f/2 camera lens. Camera lenses face similar challenges as our scan lens. They have to focus distant objects, at different angles, into the same flat photographic emulsion, while faithfully depicting true angular distances with minimal aberrations. Think for example about stars on the sky, whose light arrives as beams at different angles at the camera lens. Addressing this problem lead to lens designs that were very similar to our scan lens. One important design, sometimes referred to as a Gauss doublet objective \cite{Kidger2001}, features two symmetrically configured doublets providing much of the optical power, while additional lenses control aberrations. For example, the Zeiss Planar lens from 1897 features 6 elements in 4 groups with a symmetric pair of doublets in a design very similar to our scan lens assembly \cite{Rudolf1897}. To help students and researchers explore these sometime challenging subjects theoretically, we provide all Zemax files for the microscope optics. More advanced optimization in Zemax requires the use of their programming API. For example, the simulated point spread functions and their analysis in Fig.~\ref{fig2} were computed in Matlab, using the Zemax API. The code is provided in our repository. This allows for fast optimization of system parameters by effectively scanning through lens spacing, lens types, and the like. This can allow researchers and students to explore the limits and possible future improvements of the system. Experimental modifications of the system are simpler in the format of a workshop. For example, students can swap or rotate lenses and observe the effects. The scan lens assembly is particularly sensitive to this, because the incoming laser beam is concentrated into a small region that gets scanned across the surface of the lens. Small surface imperfections will introduce optical difference across spot positions, and significantly affect field flatness, cf. Fig.~\ref{fig5}C,D,E. \textcolor{blue}{Small misalignment in the optical path can also lead to characteristic image artifacts, for illustrations see\cite{Corbett2014, Lees2024}, that can help to develop an intuition about how the system operates.}\newline
Finally, it can be a useful learning experience to re-use parts from other microscopes, such as xy scanners from a confocal microscope\cite{Nikolenko2013}, and to mix and match parts to learn what works and what does not. It is an important learning experience to understand failure modes. For example, assembling the lens groups can lead to dust when tightening the retaining rings through the anodized aluminium threads. Or the rectangular ``TV screen'' field of view (visible in the figures at zoom level 1 above) is caused by a round, and not elliptical mirror above the scan engine. This mirror clips the beam along one axis. (This is corrected in the CAD files.) When analyzing data, we suggest to use simple programs. Our image analyses here were done in Fiji \cite{Schindelin2019} combined with simple python and matlab scripts. In our experience, students learn very effectively through hands-on open-ended pedagogy and pursuing their own interests \cite{Zajdel2016}.\newline
\subsection*{Open scientific instrumentation}
-In this article, we described the instrument's mode of operation, and provide an overview of the design. For the complete and illustrated built instructions, please see \nameref{S1_Appendix} and our \nameref{repository}. To aid discoverability, we have generated an Open Know How Manifest (OKH-manifest), located in the Online Repository. Our project is also certified by the open source hardware association (OSHWA UID US002677).\newline
-The growing space of open-source instrumentation can improve science in several ways: (1) enabling faster innovation through lower costs and enabling work in low-resource settings. (2) facilitating review and inspection by avoiding black-box designs and (3) providing a benchmark for innovation towards next-generation technology. We hope that our instrument, combined with earlier designs \cite{Rosenegger2014, Mayrhofer2015, Tan1999}, and recent developments like the head mounted microscope \cite{Zong2022} or the Janelia Research Campus microscope \cite{Janelia2024}, can serve as a benchmark for future instruments. Relatedly, we have contributed to other projects that follow this philosophy in microscopy \cite{Scott2018}, but also designing an open source ventilator during the COVID-19 pandemic \cite{LaChance2022,POVMC2022}, and a \textmu-contact printing device \cite{Samhaber2016}. Making these devices fully open has informed the narrative in this article, the design and organization of the published files, and also the choice of an open access journal. Finally, the open approach can make two-photon microscopy more accessible to researchers, thereby greatly expanding the ability to train students in approaching such technology and producing new inventions. In our ventilator work, this has led to one of the first devices using deep reinforcement learning that performed superior to classical control schemes \cite{Suo2021}. Future will tell how the community will make best use of Twinkle. The world is moving towards more open technology \cite{Schottdorf2024}, in particular in the microscopy community \cite{Hohlbein2022}. We hope that our microscope and its future iterations can play a part in this movement.
+In this article, we described the instrument's mode of operation, and provide an overview of the design. For complete and illustrated built instructions, see the attached \nameref{S1_Appendix} and our \nameref{repository}. To aid discoverability, we have generated an Open Know How Manifest (OKH-manifest), located in the Online Repository. Our project is also certified by the open source hardware association (OSHWA UID US002677).\newline
+The growing space of open-source instrumentation can improve science in several ways: (1) enabling faster innovation through lower costs and enabling work in low-resource settings. (2) Facilitating review and inspection by avoiding black-box designs and (3) providing a benchmark for innovation towards next-generation technology \cite{Lees2024}. We hope that our instrument, combined with earlier designs \cite{Rosenegger2014, Mayrhofer2015, Tan1999}, and recent developments, like the head mounted microscope \cite{Zong2022}, or the Janelia Research Campus microscope \cite{Janelia2024}, can serve as a benchmark for future instruments. Relatedly, we have contributed to other projects that follow this philosophy in microscopy \cite{Scott2018}, but also designing an open source ventilator during the COVID-19 pandemic \cite{LaChance2022,POVMC2022}, and a \textmu-contact printing device \cite{Samhaber2016}. Making these devices fully open has informed the narrative in this article, the design and organization of the published files, and also the choice of an open access journal. Finally, the open approach can make two-photon microscopy more accessible to researchers, thereby greatly expanding the ability to train students in approaching such technology and producing new inventions. In our ventilator work, this has led to one of the first devices using deep reinforcement learning that performed superior to classical control schemes \cite{Suo2021}. Future will tell how the community will make best use of Twinkle. The world is moving towards more open technology \cite{Schottdorf2024}, in particular in the microscopy community \cite{Hohlbein2022}. We hope that our microscope and its future iterations can play a part in this movement.
\subsection*{Disclaimer}
-The aim of our work is to allow anyone to build a high-performance and cost-effective two-photon microscope. However, the information here cannot be exhaustive. We recommend anyone embarking on this journey to engage with the community, to carefully think about effective solutions to their use-case and specific experiment at hand, and to adjust to the ever changing landscape of available technology. When adapting the optical path to their needs, we suggest to start at our lens combinations, and then optimize (at the very least) the lens spacings in Zemax. Technology develops rapidly, and many of the specific components mentioned here will be outdated in a few years. We believe that science progresses best if researchers understand their instrumentation, and leverage the current cutting edge of what is possible to further their science.\newline
+The aim of our work is to allow anyone to build a high-performance and cost-effective two-photon microscope. However, the information here cannot be exhaustive. We recommend anyone embarking on this journey to engage with the community, to carefully think about effective solutions to their use-case and specific experiment at hand, and to adjust to the ever changing landscape of available technology. When adapting the optical path to their needs, we suggest starting at our lens combinations and then optimizing (at the very least) the lens spacings in Zemax. Technology develops rapidly and many of the specific components mentioned here will be outdated in a few years. We believe that science progresses best if researchers understand their instrumentation and leverage the current cutting edge of what is possible to further their science.\newline
This work is free: you can redistribute it and/or modify it under the terms of the Creative Commons Attribution 4.0 International license, version 4 of the License, or (at your option) any later version (CC-BY-4.0). This work is distributed in the hope that it will be useful, but without any warranty, to the extent permitted by law; without even the implied warranty of merchantability or fitness for a particular purpose. A copy of the License is provided in our repository. For more details, see \url{http://www.gnu.org/licenses/}.
\section*{Supporting information}
-\paragraph*{S1 Appendix}
+\paragraph*{S1 Appendix. This contains detailed construction instructions and technical details.}
\label{S1_Appendix}
-A document with detailed build instructions and pictures of the microscope.
+\textcolor{blue}{A \texttt{.pdf} file with numerous pictures, annotations, and detailed step-by-step building instructions for the microscope. The raw code, pictures, and vectorized versions of the figures are available in the online repository.}
-\paragraph*{Online Repository}
+\paragraph*{Online Repository. This contains all the underlying materials of this article.}
\label{repository}
-All CAD and optics design files, bill of materials, and other useful information: \url{https://github.com/BrainCOGS/Microscope}.
+\textcolor{blue}{A repository, hosted on github, including code, \texttt{.svg} figures, and the raw \texttt{.tex} files for this article and its supplement. In addition, we provide all CAD and optical design files, material billing and other useful information. The CAD files are provided in several standard formats. We have used Fusion 360. The optical design was performed in Ansys' OpticStudio (Zemax): \url{https://github.com/BrainCOGS/Microscope}.}
\section*{Acknowledgments}
-MS and MD are supported by NIH grant U19NS132720. MS is also supported by a C.V. Starr fellowship and a Burroughs Wellcome Fund's Career Award at the Scientific Interface. AL was supported by the NSF through the Center for the Physics of Biological Function (PHY-1734030). We thank Beatrice Hadiwidjaja for transgenic Zebrafish husbandry, and Drs. Lindsay Collins and Anthony Ambrosini for hospitality and accommodating our tinkering in the teaching lab of the Princeton Neuroscience Institute. We thank Kai Br\"oking for helpful comments.
+\textcolor{blue}{We thank Beatrice Hadiwidjaja for the transgenic zebrafish husbandry and Drs. Lindsay Collins and Anthony Ambrosini for hospitality and accommodating our tinkering in the teaching lab of the Princeton Neuroscience Institute. We thank Kai Br\"oking and Peter Rupprecht for helpful comments on the manuscript.}
\section*{Author Contributions}
% Follow the "CRediT Taxonomy" - plos standard.
\begin{itemize}
\item \textbf{Conceptualization}: Stephan Y. Thiberge.
- \item \textbf{Data curation}: Manuel Schottdorf, Mika Diamanti, Albert Lin, Stephan Y. Thiberge.
+ \item \textbf{Data curation}: Manuel Schottdorf, E. Mika Diamanti, Albert Lin, Stephan Y. Thiberge.
\item \textbf{Formal analysis}: Manuel Schottdorf, E. Mika Diamanti, Albert Lin.
\item \textbf{Funding acquisition}: Stephan Y. Thiberge.
- \item \textbf{Investigation}: Manuel Schottdorf, P. Dylan Rich, E. Mika Diamanti, Albert Lin, Sina Tafazoli, Edward Nieh.
+ \item \textbf{Investigation}: Manuel Schottdorf, P. Dylan Rich, E. Mika Diamanti, Albert Lin, Sina Tafazoli, Edward H. Nieh.
\item \textbf{Methodology}: Manuel Schottdorf, P. Dylan Rich, E. Mika Diamanti, Albert Lin, Sina Tafazoli, Edward H. Nieh, Stephan Y. Thiberge.
\item \textbf{Project administration}: Manuel Schottdorf, Stephan Y. Thiberge.
\item \textbf{Resources}: Manuel Schottdorf, Stephan Y. Thiberge.
@@ -302,7 +308,7 @@ \section*{Author Contributions}
\item \textbf{Validation}: Manuel Schottdorf, P. Dylan Rich, E. Mika Diamanti.
\item \textbf{Visualization}: Manuel Schottdorf, E. Mika Diamanti, Albert Lin.
\item \textbf{Writing – original draft}: Manuel Schottdorf.
- \item \textbf{Writing – review \& editing}: Manuel Schottdorf, P. Dylan Rich, E. Mika Diamanti, Albert Lin, Sina Tafazoli, Stephan Thiberge.
+ \item \textbf{Writing – review \& editing}: Manuel Schottdorf, P. Dylan Rich, E. Mika Diamanti, Albert Lin, Sina Tafazoli, Stephan Y. Thiberge.
\end{itemize}
\nolinenumbers
@@ -315,15 +321,129 @@ \section*{Author Contributions}
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+Peter Rupprecht. (2019). PTRRupprecht/Photon-yield-and-pulse-dispersion: Photon yield and pulse dispersion, version 1 (Version version1). Zenodo. https://doi.org/10.5281/zenodo.2592465 \url{https://ptrrupprecht.github.io/Photon-yield-and-pulse-dispersion/}.
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+Smith, SL (2019). Building a Two-Photon Microscope Is Easy. In: Hartveit, E. (eds) Multiphoton Microscopy. Neuromethods, vol 148. Humana, New York, NY. \url{https://doi.org/10.1007/978-1-4939-9702-2_1}.
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@@ -526,21 +552,36 @@ \section*{Author Contributions}
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-Zajdel TJ, Maharbiz MM (2016) Teaching design with a tinkering-driven robot hack. Proceedings of 2016 IEEE Frontiers in Education Conference (FIE) \url{https://ieeexplore.ieee.org/document/7757484}
+Zajdel TJ, Maharbiz MM (2016) Teaching design with a tinkering-driven robot hack. Proceedings of 2016 IEEE Frontiers in Education Conference (FIE) \url{https://ieeexplore.ieee.org/document/7757484}.
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@@ -552,16 +593,16 @@ \section*{Author Contributions}
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\bibitem{POVMC2022}
-The Princeton Open Ventilation Monitor Collaboration et al. Inexpensive Multipatient Respiratory Monitoring Syshttps://www.yapsutube.com/watch?v=uYOF-YggWAgtem for Helmet Ventilation During COVID-19 Pandemic. ASME. J. Med. Devices. March 2022; 16(1): 011003. https://doi.org/10.1115/1.4053386
+The Princeton Open Ventilation Monitor Collaboration et al. Inexpensive Multipatient Respiratory Monitoring System for Helmet Ventilation During COVID-19 Pandemic. ASME. J. Med. Devices. 2022 March; 16(1):011003. doi: 10.1115/1.4053386.
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Samhaber R, Schottdorf M, El Hady A, Bröking K, Daus A, Thielemann C, Stühmer W, Wolf F. Growing neuronal islands on multi-electrode arrays using an accurate positioning-\textmu CP device. J Neurosci Methods. 2016 Jan 15;257:194-203. doi: 10.1016/j.jneumeth.2015.09.022. Epub 2015 Oct 1. PMID: 26432934.
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+Suo D, Ghai U, Minasyan E, Gradu P, Chen X, Agarwal N, Zhang C, Singh K, LaChance J, Zajdel T, Schottdorf M, Cohen D, Hazan E. Machine Learning for Medical Ventilator Control. Machine Learning for Health (ML4H) at NeurIPS. 2021. doi: 10.48550/arXiv.2111.10434. arXiv: 2102.06779 (2022)
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+
+
+
-
+ x="57.069496"
+ y="836.72528"
+ clip-path="url(#clipPath70514)"
+ transform="translate(0.29096415,-1.5699397)" />
-
+ y="838.29028"
+ clip-path="url(#clipPath2550)"
+ transform="translate(-5.7015605,-3.9478948)" />
SFig. 11
+ y="864.95679">SFig. 11
A
+ style="font-style:normal;font-variant:normal;font-weight:bold;font-stretch:normal;font-size:4.23333px;font-family:'Liberation Sans';-inkscape-font-specification:'Liberation Sans Bold';font-variant-ligatures:normal;font-variant-caps:normal;font-variant-numeric:normal;font-variant-east-asian:normal;fill:#ffffff;fill-opacity:1;stroke-width:0.265;stroke-miterlimit:4;stroke-dasharray:none"
+ x="0.66308117"
+ y="838.70264">A
B
+ style="font-style:normal;font-variant:normal;font-weight:bold;font-stretch:normal;font-size:4.23333px;font-family:'Liberation Sans';-inkscape-font-specification:'Liberation Sans Bold';font-variant-ligatures:normal;font-variant-caps:normal;font-variant-numeric:normal;font-variant-east-asian:normal;fill:#ffffff;fill-opacity:1;stroke-width:0.265;stroke-miterlimit:4;stroke-dasharray:none"
+ x="57.760647"
+ y="838.70264">B
+ C
C
B
+ x="7.0000415"
+ y="861.07782">B
B
+ x="107.35502"
+ y="839.61926">B
C
+ x="107.22729"
+ y="844.06989">C
D
+ x="107.22729"
+ y="847.67743">D
A
+ x="95.287743"
+ y="840.95911">A
E
+ x="95.323723"
+ y="847.75543">E
H
+ x="101.67722"
+ y="852.76501">H
F
+ x="88.819489"
+ y="854.06">F
O
- O
+ x="112.2539"
+ y="870.84045">O
N
- N
+ x="108.54563"
+ y="865.45294">N
M
+ x="101.90678"
+ y="866.79382">M
M
- I
+ x="109.16985"
+ y="854.33752">I
G
+ x="96.959648"
+ y="854.89252">G
J
+ x="63.237049"
+ y="848.03149">J
K
+ x="66.989136"
+ y="853.04248">K
L
+ x="69.394173"
+ y="853.04248">L
A
+ x="15.415804"
+ y="854.74335">A
C
+ x="28.201437"
+ y="864.25232">C
D
+ x="37.001415"
+ y="837.93811">D
F
+ x="39.703056"
+ y="867.71466">F
to Control
-
+ x="115.98096"
+ y="862.10101" />
-
12 VPower supply
+ style="fill:#f9f9f9;fill-opacity:1;stroke-width:0.264583"
+ x="58.434383"
+ y="845.15814"
+ id="tspan36314">12 V supply
±28 VPower supply
- Inside view
+ style="fill:#fefefe;fill-opacity:1;stroke-width:0.264583"
+ x="86.735443"
+ y="837.54169"
+ id="tspan36314-0">±28 V supply
-
- Resonant
- Slow
-
13 μm
+ +
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+ +
+
+
+ +28V
+ -28V
+ 0V
+ F
+ G
+ H
+ I
+
+
+
+ O
+ N
+ M
+
+ Inside view
+ Resonant
+ Slow
+ D
+ E
+
diff --git a/documentation/supplement.tex b/documentation/supplement.tex
index ea7a32d..5e88379 100644
--- a/documentation/supplement.tex
+++ b/documentation/supplement.tex
@@ -26,6 +26,7 @@
\usepackage{enumerate}
\usepackage{sidecap}
\usepackage{wasysym}
+\usepackage{mathbbol}
% create \thickcline for thick horizontal lines of variable length
\newlength\savedwidth
@@ -69,6 +70,7 @@
%% Include all macros below
\newcommand{\microns}{~\textmu m~} % ALL UNITS UPRIGHT WITH SPACE
\newcommand{\invivo}{\textit{in vivo}~}
+\newcommand{\complexi}{i}
%% END MACROS SECTION
\begin{document}
@@ -91,15 +93,16 @@ \subsection{Safety}
Class 4 lasers are extremely dangerous. A reflection on a surface can be enough to inflict permanent eye damage. Safety protocols must be observed. Consult with your institution's environmental health and safety officer and take proper training before performing any work with a laser system. Learn from people experienced with these devices and work carefully.\newline
During alignment, always wear goggles (ours were ML1 by NoIR LaserShields with an optical density $>7$ in the relevant range around $920\text{ nm}$), remove jewelry and watches, and use the low laser power (“alignment mode”) whenever possible. Once the assembly progresses further, full power mode is necessary. Extra care must be taken then. Always use Laser Viewing Cards to track the laser. We use Thorlabs VRC4. Always be conscious where the beam is, and where the beam goes. The beam must be terminated in beam blockers and should be shielded whenever possible with opaque plastic or anodized aluminium tubes to increase safety.
-\subsection{Miscellaneous tools}
+\subsection{Miscellaneous tools and tips}
+The following tools are used throughout the build process:
\begin{itemize}
- \item Use a high quality duster to remove any residue from optical surfaces. We use a Miller-Stephenson Aero-Duster Ultra MS-222L, in addition to lens tissues (MC-5, Thorlabs) and Methanol (Sigma-Aldrich).
+ \item A high quality duster to remove any residue from optical surfaces. We use a Miller-Stephenson Aero-Duster Ultra MS-222L, in addition to lens tissues (MC-5, Thorlabs) and Methanol (Sigma-Aldrich).
\item To glue optical filters we used an extra fast setting epoxy (Hardman Double/Bubble Epoxy). To seal small gaps, we use black silicone putty (V-tie self-setting silicone rubber).
\item We used either McMaster or Thorlabs screwdrivers and hex keys.
\item For alignment, we used laser viewing cards (VRC4, Thorlabs), cage system alignment plates (VRC4CPT, Thorlabs), and irises (SM1 Graduated Ring-Actuated Iris Diaphragm, SM1D12C, Thorlabs).
\end{itemize}
-
-\subsection{Miscellaneous tips}
+%
+The following general tips might be of help as well:
\begin{itemize}
\item Shield the beam as soon as alignment of a particular section is done. This increases safety, and avoids accidentally bumping into aligned components.
\item \textit{Walking the beam}: Any beam can be aligned through two irises using two mirrors. The first mirror the beam meets is sometimes called the ``position'' mirror as it affects primarily the position of the beam. The second ``angle'' mirror instead primarily affects the angle. Assigning each mirror a separate function (position versus angle) and by adjusting them iteratively, is a good rule of thumb for driving the beam through the 2 irises and eventually hitting a specific point at the desired angle \cite{Thorlabs}.
@@ -146,11 +149,11 @@ \subsection{Structure and use of this document}
\section{Assembly on the optical table}
%
\subsection{Beam conditioning}
-An overview of the system for beam conditioning is shown in Fig.~\ref{sfig1} with key items labelled by capital letters. The following lines will cover the items, and their alignment.
+An overview of the system for beam conditioning is shown in Fig.~\ref{sfig1} with key items labeled by capital letters. The following lines will cover the items, and their alignment.
%
\begin{figure}[!t]
\includegraphics[width=\textwidth]{sfig1.jpg}
- \caption{{\bf Overall view of the beam conditioning path on the optical table.} Shown is a photograph of the components used to condition the beam for our microscope. Key components of the system are labelled with capital letters. The laser is shown as a red transparent line with intensity color coded. The beamsplitter cube (component D) splits the beam into two, halving its power. The Pockels cell (component G) reduces the power further. The reflections out of the Pockels cell and from the shutter (item J) are shown in yellow. The telescope (item K) magnifies the beam $\times 2$. The empty SM1 tube is an anodized aluminum SM1 lens tube cover (SC600, Thorlabs). All parts are explained in the text together with a note on their alignment.}
+ \caption{{\bf Overall view of the beam conditioning path on the optical table.} Shown is a photograph of the components used to condition the beam for our microscope. Key components of the system are labeled with capital letters. The laser is shown as a red transparent line with intensity color coded. The beamsplitter cube (component D) splits the beam into two, halving its power. The Pockels cell (component G) reduces the power further. The reflections out of the Pockels cell and from the shutter (item J) are shown in yellow. The telescope (item K) magnifies the beam $\times 2$. The empty SM1 tube is an anodized aluminum SM1 lens tube cover (SC600, Thorlabs). All parts are explained in the text together with a note on their alignment.}
\label{sfig1}
\end{figure}
%
@@ -218,7 +221,7 @@ \subsubsection{Components}
%
\begin{figure}[h]
\includegraphics[width=\textwidth]{sfig4.jpg}
- \caption{{\bf Overall view of the scanning post.} \textbf{A)} Side few. Key components of the system are labelled with capital letters. These are explained in the text together with a note on how to align these components. Importantly, the machined block (C) houses the scanning mirrors, and the scan lens assembly is placed in the SM2 house labelled (E). \textbf{B)} Frontal view. The PMT preamplifiers (component G) and the scanning mirror power supply (component H) are visible.}
+ \caption{{\bf Overall view of the scanning post.} \textbf{A)} Side few. Key components of the system are labeled with capital letters. These are explained in the text together with a note on how to align these components. Importantly, the machined block (C) houses the scanning mirrors, and the scan lens assembly is placed in the SM2 house labeled (E). \textbf{B)} Frontal view. The PMT preamplifiers (component G) and the scanning mirror power supply (component H) are visible.}
\label{sfig4}
\end{figure}
%
@@ -250,7 +253,7 @@ \subsubsection{Components}
%
\begin{figure}[t!]
\includegraphics[width=\textwidth]{sfig5.jpg}
- \caption{{\bf Overall view of the microscope head.} Key components of the system are labelled with capital letters. These are explained in the text together with a note on how to align these components.}
+ \caption{{\bf Overall view of the microscope head.} Key components of the system are labeled with capital letters. These are explained in the text together with a note on how to align these components.}
\label{sfig5}
\end{figure}
%
@@ -292,7 +295,7 @@ \subsubsection{Components}
%
\begin{SCfigure}
\includegraphics[width=.5\textwidth]{sfig6.jpg}
- \caption{{\bf Side view of the microscope head.} Key components of the system are labelled with capital letters. These are explained in the text together with a note on how to align these components.\newline
+ \caption{{\bf Side view of the microscope head.} Key components of the system are labeled with capital letters. These are explained in the text together with a note on how to align these components.\newline
Set up like this, we were measuring the amount of light traveling through the system. Note the detector taped to the objective with green tape.\newline
Also note that the photomultiplier tubes (component C) are light sensitive and need to be installed in the dark. We recommend to switch off the room lights when handling them, and allowing your eyes to adapt to the dark over several minutes. After installation into the collection optics, the lights can safely be used. The objective is easy to install, and just screwed into the piezoelectric collar, component F.}
\label{sfig6}
@@ -387,11 +390,11 @@ \subsection{Scan lens assembly}
\subsection{Assembly of the collection optics}
%
-The collection optics are made from several components, including optical filters, lenses, and custom metal pieces. Fig.~\ref{sfig9}A shows a cut through the CAD drawing. The function is described in the main manuscript. A pictures of the collection optics box is shown in Fig.~\ref{sfig9}B with key components labelled. Here, we will focus on the assembly and their alignment.
+The collection optics are made from several components, including optical filters, lenses, and custom metal pieces. Fig.~\ref{sfig9}A shows a cut through the CAD drawing. The function is described in the main manuscript. A pictures of the collection optics box is shown in Fig.~\ref{sfig9}B with key components labeled. Here, we will focus on the assembly and their alignment.
%
\begin{figure}[h]
\includegraphics[width=\textwidth]{sfig9.jpg}
- \caption{{\bf Assembly of the collection optics.} \textbf{A)} The assembled collection optics with bottom cover removed, perspective from below. \textbf{B)} The collection optics during the assembly process, similar to panel A. Key components are labelled (see text). EDITS TO MAKE IN THE ARROWS}
+ \caption{{\bf Assembly of the collection optics.} \textbf{A)} The assembled collection optics with bottom cover removed, perspective from below. \textbf{B)} The collection optics during the assembly process, similar to panel A. Key components are labeled (see text). EDITS TO MAKE IN THE ARROWS}
\label{sfig9}
\end{figure}
%
@@ -419,7 +422,7 @@ \subsection{Assembly of the scanning mirror housing}
%
\begin{figure}[!t]
\includegraphics[width=\textwidth]{sfig10.jpg}
- \caption{{\bf The scanning mirror box.} \textbf{A)} The custom aluminium box housing the pair of scanning mirrors. \textbf{B)} Close-up of the same box with key parts labelled: A: Two windows (WG11010-B, Thorlabs) to keep dust out. B: Attachment of the box to the 95 mm optical rail. C: An Iris used for alignment, and D: SM1 tubing. \textbf{C)} Bottom part of the aluminium box. A: the windows, and B: screws to attach the box to the drop-on rail carriage via four 0.75 inch tall 1 inch diameter posts. \textbf{D)} The scanning mirrors are mounted to the lid using heat transfer compound. E: Anodized aluminium block holding the scanning mirrors. F: One of the scanning mirror assemblies, G: The aluminium lid and H: The scanning mirrors. \textbf{E)} Same as \textbf{D} but different angle. \textbf{F)} Same as \textbf{E} but different angle. I: Cables controlling the scanning mirrors.}
+ \caption{{\bf The scanning mirror box.} \textbf{A)} The custom aluminium box housing the pair of scanning mirrors. \textbf{B)} Close-up of the same box with key parts labeled: A: Two windows (WG11010-B, Thorlabs) to keep dust out. B: Attachment of the box to the 95 mm optical rail. C: An Iris used for alignment, and D: SM1 tubing. \textbf{C)} Bottom part of the aluminium box. A: the windows, and B: screws to attach the box to the drop-on rail carriage via four 0.75 inch tall 1 inch diameter posts. \textbf{D)} The scanning mirrors are mounted to the lid using heat transfer compound. E: Anodized aluminium block holding the scanning mirrors. F: One of the scanning mirror assemblies, G: The aluminium lid and H: The scanning mirrors. \textbf{E)} Same as \textbf{D} but different angle. \textbf{F)} Same as \textbf{E} but different angle. I: Cables controlling the scanning mirrors.}
\label{sfig10}
\end{figure}
%
@@ -429,23 +432,24 @@ \subsection{Assembly of the scanning mirror housing}
\subsection{Scanning mirror power supply}
%
-The scanning mirrors come with driver circuit boards. We place them in a prototype box (EG6, Gold Box, Acopian), and connect the driver board's power supply and signal lines to connectors for easier access, see Fig.~\ref{sfig11}A. The driver boards are mounted to the aluminium walls of the prototype box with heat transfer compound (Wakefield Thermal Joint Compound Type 120-5 Silicone). The power connectors accept banana plugs. The signals are connected to BNC connectors.
+The scanning mirrors come with driver circuit boards. We place them in a prototype box (EG6, Gold Box, Acopian), and connect the driver board's power supply and signal lines to connectors for easier access, see Fig.~\ref{sfig11}A. The driver boards are mounted to the aluminium walls of the prototype box with heat transfer compound (Wakefield Thermal Joint Compound Type 120-5 Silicone). The power connectors accept banana plugs. The signals are connected to BNC connectors.
%
\begin{figure}[t]
\includegraphics[width=\textwidth]{sfig11.jpg}
- \caption{{\bf Scanner electronics.} \textbf{A)} Overview of all scanning mirror parts. Key parts are labelled with capital letters. Optomechanics were covered in Fig.~\ref{sfig10}. All electrical drivers were placed in the yellow circuit box. \textbf{B)} Power supply and connection of the scanning mirrors with key parts labelled by capital letters. The inset shows the scanning mirror driver boards provided by Novanta and mounted in the yellow circuit box. The smaller driver circuit (blue) powers the resonant scanner. The larger circuit board (green) drives the slow galvanometer. The circuit boards are mounted to the aluminium walls with screws and heat transfer compound.}
+ \caption{{\bf Scanner electronics.} \textbf{A)} Overview of all scanning mirror parts. Key parts are labeled with capital letters. Optomechanics were covered in Fig.~\ref{sfig10}. All electrical drivers were placed in the yellow circuit box. \textbf{B)} Power supply and connection of the scanning mirrors with key parts labeled by capital letters. \textcolor{blue}{\textbf{C)} Wiring diagram for the $\pm 28\text{ V}$ power supply, using two galvanically isolated supplies. \textbf{D)} The scanning mirror driver boards, provided by Novanta, and mounted in the yellow circuit box. The smaller driver circuit (blue) powers the resonant scanner. The larger circuit board (green) drives the slow galvanometer. The circuit boards are mounted to the aluminium walls with screws and heat transfer compound. \textbf{E)} Same box as in D, but from the outside and key parts labeled by capital letters.}}
\label{sfig11}
\end{figure}
%
-Fig.~\ref{sfig11}A shows as the housing box (component A) and xy scanning mirror set (component B) that we discussed above. Components C are the cables connecting the scanning mirrors to their driver boards. These driver boards are housed in a yellow prototype box, and connected to accept power (component D), and signals (component F). Fig.~\ref{sfig11}B shows the power supply of the scanning mirrors. At time of purchase, we requested the controller of the slow galvanometer mirror to be configured for usage at $\pm 28\text{ V}$ (manufacturer default is $\pm 24\text{ V}$). This overall improves dynamical properties. The controller requires three voltage inputs, respectively -28 V, 0 V, and +28 V. The 0 V is also grounded. The second power supply to the left provides 12 V for the resonant scanner. The supplies are connected to the power supply ports on the prototype box, and the signal lines are connected to the MBF Bioscience DAQ system. Key components in the figure are labelled with capital letters. These are as follows:
+Fig.~\ref{sfig11}A shows as the housing box (component A) and xy scanning mirror set (component B) that we discussed above. Components C are the cables connecting the scanning mirrors to their driver boards. These driver boards are housed in a yellow prototype box, and connected to accept power (component D), and signals (component F). Fig.~\ref{sfig11}B shows the power supply of the scanning mirrors. At time of purchase, we requested the controller of the slow galvanometer mirror to be configured for usage at $\pm 28\text{ V}$ (manufacturer default is $\pm 24\text{ V}$). This overall improves dynamical properties. The controller for the slow scanner requires three supplies respectively: -28 V, 0 V, and +28 V. \\
+\textcolor{blue}{To provide these three signals, we used a power supply with two galvanically isolated and adjustable output channels, that can be connected in series (Note: not every power supply can do this. We used a GPD-3303D from Gw Instek that allows this function with the SER/INDEP key). The voltage of each of the two unconnected channels was set to +28 V. To produce the three supply signals, we used these two channels in series. In other words, we connect the positive terminal of the first channel to the negative terminal of the second channel. As the channels are otherwise not connected to a common ground (this is meant with the words ``galvanically isolated''), this does not produce a short circuit. Instead, this link effectively provides a new reference point, relative to which the first supply produces a negative -28 V supply, and the second supply produces the +28 V supply. This is illustrated in Fig.~\ref{sfig11}C. In this arrangement, a multimeter would read between points G and I +28 V, between G and F -28 V, and between F and I +56 V. The supply of the resonant scanners is simpler, as it only requires a single +12 V line. The supplies are connected to the power supply ports on the prototype box, in which the two driver boards reside, see Fig.~\ref{sfig11}D. Signal lines are connected on the outside with coaxial cables to the MBF Bioscience DAQ system, see Fig.~\ref{sfig11}E.} Key components in the panels are labeled with capital letters. These are as follows:
\begin{itemize}
\item \textbf{A)} Power supply (GPD-3303D, Gw Instek) with two channels configured in series, both set to 28 V.
\item \textbf{B)} Have CH 1 turned on.
- \item \textbf{C)} Press the ``Ser/Indep'' button. This wires the two channels together to a shared references to provide the $\pm 28\text{ V}$ and the 0 V reference line.
- \item \textbf{D)} This enables our power supply unit. Only when pressed, power is sourced to the galvanometer driver board.
+ \item \textbf{C)} Press the ``Ser/Indep'' button. This wires the two channels together to a shared references to provide the $\pm 28\text{ V}$ and the 0 V reference line, see Fig.~\ref{sfig11}C.
+ \item \textbf{D)} This button enables our power supply unit. Only when pressed, power is sourced to the galvanometer driver board.
\item \textbf{E)} CC/CV indicator light. Make sure that the device is set to CV (constant voltage).
- \item \textbf{F)} The -28 V output line (black). This is connected to the -28 V line on the electronics box (shown below).
- \item \textbf{G)} The reference lines, 0 V (white) connected to the 0 V line on the electronics box.
+ \item \textbf{F)} The -28 V output line (black). This is connected to the -28 V line on the electronics box (shown below in Fig.~\ref{sfig11}E).
+ \item \textbf{G)} The reference lines, 0 V (white) connected to the 0 V line on the electronics box, see Fig.~\ref{sfig11}E.
\item \textbf{H)} Connects the reference line to ground.
\item \textbf{I)} The +28 V output line (red) connected to the +28 V line on the electronics box. Conform the correct voltage with a reliable multimeter BEFORE powering up the galvanometer controller, e.g. with a Fluke 117.
\item \textbf{J)} Power supply (E3630A, Keysight) set to 12 V to power the resonant scanners.
@@ -462,7 +466,7 @@ \subsection{Assembly of PMT control circuits}
%
\begin{figure}[t]
\includegraphics[width=\textwidth]{sfig12.jpg}
- \caption{{\bf Custom electronics.} \textbf{A)} Picture of the control box. Component A are the displays for the control voltage for red and green channels. Component B are the two potentiometers to set the two channel gains. \textbf{B)} Inside view of the potentiometer box with key components labelled (see text). \textbf{C)} Connector attached to the controller box before plugging into the PMTs. \textbf{D)} Connector from the Control box to the PMT cables. \textbf{E)} Outputs of the PMTs. Note the four color-coded American wire gauge \#28 cables, and the coaxial signal line\cite{Hamamatsu2024}.}
+ \caption{{\bf Custom electronics.} \textbf{A)} Picture of the control box. Component A are the displays for the control voltage for red and green channels. Component B are the two potentiometers to set the two channel gains. \textbf{B)} Inside view of the potentiometer box with key components labeled (see text). \textbf{C)} Connector attached to the controller box before plugging into the PMTs. \textbf{D)} Connector from the Control box to the PMT cables. \textbf{E)} Outputs of the PMTs. Note the four color-coded American wire gauge \#28 cables, and the coaxial signal line\cite{Hamamatsu2024}.}
\label{sfig12}
\end{figure}
%
@@ -628,12 +632,51 @@ \section{Recommendations for use}
\item Measure (1) the laser's power levels, (2) the Pockels cell calibration and (3) check exposed lenses and mirrors for dust and fingerprints with a flashlight. This is fast and should be done before every experiment.
\item When the microscope is not in use, it is good practice to switch off all components, except the scanning mirror power supplies, the PMT preamplifiers, and the photodiode.
\item Every once in a while, switch the red and green PMTs to check for signs of aging and validate their performance on a fluorescent slide or sample of known brightness. The PMTs can degrade and may need to be replaced, depending on use, after several years. They should be treated with care: power ramped up and down slowly, and saturation should be avoided. We use a rotation system in which new PMTs are typically used for the green channel. When their performance becomes worse, they often have a few more months or years in the (typically brighter) red channel in them.
- \item It is recommended to periodically image a bead sample to validate the PSF, as well as a Fluorescein bath to detect possible misalignment.
+ \item It is recommended to periodically image a bead sample to validate the PSF, as well as a Fluorescein bath to detect possible misalignment. For a detailed protocol, see Lees et al. \cite{Lees2024}.
\item Be mindful of imaging artifacts. When suspecting an issue, go back to the basics, and isolate the error.
\end{itemize}
%
For safety, it is good practice to shield the laser path. In addition to safety concerns, it will also help in keeping dust and dirt away from optical components.
+\section{Deriving the intensity-dispersion relation}
+\textcolor{blue}{In the main article, we stated that the expected fluorescence $I(\varphi)$ as function of group delay dispersion $\varphi$ applied to a $\varphi_0$-dispersed Gaussian pulse of full-width-at-half-maximum $\Delta\tau$ can be written as
+\begin{eqnarray}
+ I(\varphi)=I_0 \left(1+16\log(2)^2\frac{(\varphi+\varphi_0)^2}{\Delta\tau^4}\right)^{-1/2}\label{main}.
+\end{eqnarray}
+To derive this expression, consider the electrical field of a Gaussian pulse of amplitude $E_0$, width $\Delta t$, center frequency $\omega_0$, phase $\Theta$, and centered at time $t=0$:
+\begin{eqnarray}
+E(t)=E_0\exp\left(-\frac{t^2}{4\Delta t^2}\right)\cdot\exp\left(-\complexi\left(\omega_0 t + \Theta\right)\right).
+\end{eqnarray}
+Its intensity is
+\begin{eqnarray}
+P(t) = |E(t)|^2 = |E_0|^2 \exp\left(-\frac{t^2}{2\Delta t^2}\right)\label{intens}.
+\end{eqnarray}
+In the Fourier domain, the spectral content of this wave package is:
+\begin{eqnarray}
+\tilde{E}(\omega)=E_0\cdot\sqrt{2}\Delta t\, \exp\left(-\complexi\,\Theta -\Delta t^2(\omega - \omega_0)^2\right).
+\end{eqnarray}
+In the linear regime, frequency-dependent dispersion changes the electric field to
+\begin{eqnarray}
+\tilde{E}(\omega)\to\tilde{E}(\omega)\cdot\exp\left(\complexi\,\phi(\omega-\omega_0)\right)
+\end{eqnarray}
+with phase shift $\phi(\omega-\omega_0)$. We approximate this function to quadratic order $\phi(\omega-\omega_0)\approx a + b(\omega-\omega_0) + \frac{c}{2}(\omega-\omega_0)^2$. With this approximation, we obtain the Fourier representation of the electric field of the dispersed pulse:
+\begin{eqnarray}
+\tilde{E}^\text{disp}(\omega)=\tilde{E}(\omega)\cdot\exp\left(\complexi\,\left(a + b(\omega-\omega_0) + \frac{c}{2}(\omega-\omega_0)^2\right)\right).
+\end{eqnarray}
+Coefficient $a$ causes a phase shift and coefficient $b$ a time shift. Both do not change pulse width. Computing the inverse Fourier transform to obtain $E^\text{disp}(t)$ allows to compute the intensity of the dispersed pulse as
+\begin{eqnarray}
+P^\text{disp}(t) = |E^\text{disp}(t)|^2 = |E_0|^2\frac{2\Delta t^2}{\sqrt{c^2 + 4\Delta t^4}}\exp\left(-\frac{2(b+t)^2\Delta t^2}{c^2+4\Delta t^4}\right)\label{intens_disp}.
+\end{eqnarray}
+If the average power of the laser across dispersion compensation settings $c$ is constant, the prefactor is approximately constant. Comparison with Eq. \eqref{intens} then shows that the pulse has broadened in time to $\Delta t\to \frac{\sqrt{c^2+4\Delta t^4}}{2\Delta t}$. Rewriting the widths of the dispersed pulse in the more common full-width-at-half-maximum $\Delta\tau$, using $\Delta t \to \Delta\tau/\sqrt{8\log(2)}$ for a Gaussian pulse, we obtain
+\begin{eqnarray}
+\Delta\tau^\text{disp} = \frac{\sqrt{\Delta\tau^4 + 16\log(2)^2c^2}}{2 \Delta\tau\sqrt{\log(4)}}.
+\end{eqnarray}
+The two-photon fluorescence intensity $I$ is proportional to the inverse of the pulse width, and the group delay dispersion is composed of the original dispersion $\varphi_0$ and the applied correction $\varphi$, so that $c \to \varphi_0 + \varphi$. We obtain
+\begin{eqnarray}
+I(\varphi) \propto \frac{1}{\Delta\tau^\text{disp}} = \frac{2 \Delta\tau\sqrt{\log(4)}}{\sqrt{\Delta\tau^4 + 16\log(2)^2(\varphi_0 + \varphi)^2}} = \frac{2\sqrt{\log(4)}}{\Delta\tau}\frac{1}{\sqrt{1 + 16\log(2)^2\frac{(\varphi_0 + \varphi)^2}{\Delta\tau^4}}}.
+\end{eqnarray}
+Absorbing the constants $\frac{2\sqrt{\log(4)}}{\Delta\tau}\to I_0$ yields Eq.~\eqref{main}. The same result for pulse broadening, with $16\log(2)^2\approx 7.68\dots$, is stated as Eq.~(3) in \cite{Zipfel2003}.
+}
\nolinenumbers
\begin{thebibliography}{10}
@@ -663,6 +706,12 @@ \section{Recommendations for use}
Technical note on the Photomultiplier Tube Modules Series
H16200/H16201/H16204, Hamamatsu Photonics 2014 \url{https://www.hamamatsu.com/eu/en/product/optical-sensors/pmt/pmt-module/current-output-type/H16201-40.html}
+\bibitem{Lees2024}
+Lees RM, Bianco IH, Campbell RAA, Orlova N, Peterka DS, Pichler B, Smith SL, Yatsenko D, Yu CH, Packer AM. Standardised Measurements for Monitoring and Comparing Multiphoton Microscope Systems. bioRxiv [Preprint]. 2024 Jan 23:2024.01.23.576417. doi: 10.1101/2024.01.23.576417. PMID: 38328224; PMCID: PMC10849699.
+
+\bibitem{Zipfel2003}
+Zipfel WR, Williams RM, Webb WW. Nonlinear magic: multiphoton microscopy in the biosciences. Nat Biotechnol. 2003 Nov;21(11):1369-77. doi: 10.1038/nbt899. PMID: 14595365.
+
\end{thebibliography}