Zeiss Lightsheet Z.1 Operating Manual

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Operating Manual

February 2013

ZEN 2012 (black edition)

Lightsheet Z.1

Carl Zeiss Copyright Lightsheet Z.1

Knowledge of this manual is required for the operation of the instrument. Would you therefore please make yourself familiar with the contents of this manual and pay special attention to hints concerning the safe operation of the instrument.

The specifications are subject to change; the manual is not covered by an update service.

communication of its contents are not permitted. Violations will entail an obligation to pay compensation.

Issued by Carl Zeiss Microscopy GmbH

Carl-Zeiss-Promenade 10 07745 Jena, Germany

microscopy@zeiss.com www.zeiss.com/microscopy

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INTRODUCTION Lightsheet Z.1 Carl Zeiss

Contents of this Manual

Chapter 1

Chapter 2

Chapter 3

Chapter 4

Annex

Hardware

Sample Preparation

Quick Guide

Software Operation

Lightsheet Z.1 - Overview

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CHAPTER 1 - HARDWARE Lightsheet Z.1 Content Carl Zeiss

CHAPTER 1 HARDWARE

CONTENT

Page

1 MAINTENANCE AND CLEANING ......................................................................... 3

1.1 Maintenance of the Liquid Cooling System ................................................................. 5

2 ERGODRIVE OPERATING PANEL ......................................................................... 6

3 USER INTERFACES ............................................................................................... 9

3.1 Installation and Deinstallation of the Detection Modules ........................................ 10

3.2 Adjustment – Detector Recognition ........................................................................... 15

3.2.1 Adjustment – Automatic Detector Alignment .................................................................. 15

3.2.2 Adjustment – Manual Detector Alignment ...................................................................... 17

3.2.3 Adjust the Grating Focus for the Automatic or Manual Detector Alignment Tool .............. 19

3.2.4 Cable Connections for the Detection Module "PCO.Edge" .............................................. 22

3.2.5 Cable Connections for the Detection Module "Standard" ............................................... 23

3.3 Assembly of the Sample Chamber ............................................................................. 24

3.3.1 Assembly of the Sample Chamber Windows ................................................................... 24

3.3.2 Assembly of the Sample Chamber Body and the Sample Chamber Dove Tail Slide ............ 27

3.3.3 Insertion of the Drain Connector, Luer-Lock Connectors and Blind Plugs .......................... 29

3.3.4 Insertion of Accessories for Incubation ............................................................................ 30

3.4 Removing and Inserting the Sample Chamber .......................................................... 31

3.5 Assembly of the Sample Holder ................................................................................. 32

3.5.1 Assembly of the Sample Holder for Capillary ................................................................... 32

3.5.2 Assembly of Sample Holder for Syringes .......................................................................... 34

3.6 Inserting and Removing the Sample Holder .............................................................. 35

3.7 Installation of the Incubation Modules ...................................................................... 37

3.7.1 Heating Components ..................................................................................................... 37

3.7.2 CO

3.7.3 Heating Device Humidity S1 ............................................................................................ 41

3.7.4 Registration of the incubation modules ........................................................................... 41

3.8 Switch Incubation ON and OFF ................................................................................... 43

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-Module Lightsheet Z.1 ............................................................................................. 39

CHAPTER 1 - HARDWARE Carl Zeiss Content Lightsheet Z.1

3.9 Illumination and Detection Optics ............................................................................. 43

3.9.1 Removing and Inserting the Illumination Optics Unit ....................................................... 45

3.9.2 Removing and Inserting the Detection Optics Unit........................................................... 46

3.10 Removing and Inserting the Reflector Turret for Emission Selection ...................... 47

3.11 Removing and Inserting the Reflector Turret for Laser Blocking Filter .................... 49

4 INDEX ................................................................................................................. 52

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CHAPTER 1 - HARDWARE Lightsheet Z.1 Maintenance and Cleaning Carl Zeiss

1 Maintenance and Cleaning

The maintenance to be carried out by the customer is limited to cleaning the painted surfaces, replacing glass windows and rubber seals on the sample chamber, cleaning and disinfecting the system cavity and sample chamber, temperature sensors and detection optics unit, replenishment of coolants for the incubation device and liquid cooling of the detection module as necessary, as well as the replacement of CO

sterile filters for the incubation device.

a) To clean painted surfaces proceed as follows:

• Switch the device off completely and pull the mains plug.

• Make sure that no cleaning fluid is allowed to enter the system.

• Wipe the painted surfaces with a clean cloth moistened with water to which a small amount of

cleaning agent has been added. Do not use a solvent. Dry off with a lint-free cloth.

b) To clean and disinfect the system cavity proceed as follows:

• Switch the device off completely and pull the mains plug.

• Make sure that no cleaning fluid is allowed to enter the system.

• Take care not to touch the front lens of the illumination and detection optics units. Remove these

optics as necessary.

• Remove the sample chamber for cleaning.

• Cleaning the system cavity:

Wipe the inner surfaces of the system cavity with a clean cloth moistened with water to which a small amount of cleaning agent has been added. Do not use a solvent. Dry off with a lint-free cloth.

• Disinfecting the system cavity:

Carefully wipe the inner surfaces of the sample chamber with a lint-free cloth soaked in 70 ethanol or 70

% isopropyl. Allow the solution to react for a few seconds. Repeat the procedure

once or twice as necessary. Carefully wipe off the inner surfaces with a lint-free cloth soaked in distilled water.

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c) To clean and disinfect the sample chamber proceed as follows:

• Remove the sample chamber and dismantle it into individual parts. Different procedures are required for cleaning and disinfecting the various parts of the sample chamber.

− The cover glasses must be properly disposed of in the glass waste.

− The steel parts of the chamber must be cleaned and disinfected as follows:

Rinse the parts with distilled water.

Lay the parts in an ultrasonic bath containing distilled water to which a few drops of detergent have been added.

Then rinse the steel parts thoroughly with distilled water until all detergent residue has been removed.

The parts may be laid in a disinfecting bath of 70

% ethanol (> 1 hour) and then thoroughly

rinsed with distilled water.

The steel parts of the sample chamber may also be autoclaved as necessary.

− Sealing rings, hose connectors and blank plugs must be cleaned and disinfected as follows:

These parts may not be autoclaved.

First of all lay these parts in distilled water and then in a 70 % ethanol solution for max. one hour. Rinse them thoroughly with distilled water.

d) To clean and disinfect the temperature sensor proceed as follows:

• Remove the temperature sensor from the sample chamber and loosen the connections to the main system module Lightsheet Z.1.

• Once removed, thoroughly rinse the temperature sensor with distilled water and then suspend it for one hour in a 70

% ethanol solution. Make sure that the contact plug remains dry. Complete the

cleaning process by rinsing thoroughly with distilled water.

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e) To clean and disinfect the detection optics unit proceed as follows:

• Make sure that no cleaning fluid is allowed to enter the system.

• To avoid scratches, do not wipe the front lens of the optics with dry lens paper and lens cloths.

• Remove these optics from the system cavity as necessary.

• Clean the front lens of optical elements according to general recommendations (see "The clean

microscope" www.zeiss.com/microscopy) using an optical cleaning solution (Carl Zeiss optical cleaning solution: 85

% n - hexan, 15 % isopropanol. This solution is not sold by Carl Zeiss Microscopy GmbH)

and a suitable lens cleaning paper or high purity cotton wool.

• In addition to the cleaning procedure described above, for special sterility requirements the front lens area of the detection modules can be cleaned several times with a lint-free lens cloth or lens paper soaked in 100

% ethanol. Here again, observe the general notes on cleaning the optics

("The clean microscope").

1.1 Maintenance of the Liquid Cooling System

If detection modules in your Lightsheet Z.1 are connected to a liquid cooling system, consider that the pH value of the cooling liquid changes over time.

To prevent corrosion in the cooler of the detection module, the pH value of the cooling liquid must be checked at least once a year and the liquid replaced as necessary. Observe the directions of the cooling system manufacturer in the supplied operating manual.

If you have signed a service agreement with Carl Zeiss, our service staff will perform the check as part of the maintenance procedure.

The safety data sheet for the cooling liquid should also be observed.

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CHAPTER 1 - HARDWARE Carl Zeiss ErgoDrive Operating Panel Lightsheet Z.1

2 ErgoDrive Operating Panel

1 Z-axis control 2 Rotation button (switches 6 to rotation drive) 3 Mode button (fine / coarse) 4 Axis button (switches 6 to y-axis drive) 5 X-axis control 6 Y-axis / Rotation control

Fig. 1 ErgoDrive operating panel

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CHAPTER 1 - HARDWARE Lightsheet Z.1 ErgoDrive Operating Panel Carl Zeiss

With the ErgoDrive operating panel the movement of the sample holder and therefore the sample can be controlled. It is the manual equivalent of sliders in the Specimen Navigator tool in the ZEN software (Fig. 2), with which the movement of the specimen can be controlled as well.

The function to rotate around the center of the image, Rotate around Center of image (Fig. 2) is only available through the software interface. Even if chosen there, the rotation controlled by the ErgoDrive operating panel will always rotate around the axis of the motor drive.

The steering elements are two vertical rotary controls for the x, y and rotation movement. The large rotary wheel controls the z movement (Fig. 1).

Fig. 2 ZEN software, Specimen Navigator

tool window

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The following control elements are part of the ErgoDrive operation panel:

Buttons:

The Mode button (Fig. 1/3) changes the speed of the drive from coarse to fine and back.

The Rotation button (Fig. 1/2) changes the upper rotary control to move the rotation drive (Fig. 1/6).

The Axis button (Fig. 1/4) changes the upper rotary control to move the y drive (Fig. 1/6).

Rotation control:

Upper rotary control (Fig. 1/6), after pressing the Rotation button (Fig. 1/2):

Clockwise: clockwise rotation of the sample; the angle is reduced in the software interface

Counter-clockwise: counter-clockwise rotation of the sample, the angle is increased in the

software interface

Y axis control:

Upper rotary control (Fig. 1/6), after pressing Axis button (Fig. 1/4):

Clockwise: upward movement of the sample

Counter-clockwise: downward movement of the sample

X axis control:

Lower rotary control (Fig. 1/5):

Clockwise: right movement of the sample

Counter-clockwise: left movement of the sample

Z axis control:

Large rotary wheel (Fig. 1/1):

Clockwise: sample movement towards the detection optic

Counter-clockwise: sample movement away from the detection optic

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nd lower system cavity doors and the upper

system door are equipped with a safety interlock system to guarantee laser safety. These

be manipulated. Other interfaces not described here are reserved for

ervice personnel. The following devices

Lightsheet Z.1 User Interfaces Carl Zeiss

3 User Interfaces

The ports of the detection modules, upper a

locking devices must not service and may only be used by authorized Carl Zeiss s may be mounted and dismounted by, or are accessible to, the user:

− Detection modules

− Sample chamber

− Sample Holder

− Incubation components

− Illumination optics

− Detection optics

− Reflector turret.

Customized Sample Chamber

The sample chamber delivered with the Lightsheet Z.1 can be used with a multitude of media (e.g. PBS, cell culture medium, artificial sea water) and is carefully designed to produce optimal results with the Lightsheet Z.1 system. However, if applications demand changes to the original sample chamber, Carl Zeiss Microscopy GmbH provides the CAD files and the corresponding technical drawing of our sample chamber for your convenience, in accordance with the following disclaimer:

Carl Zeiss Microscopy GmbH (hereinafter “we”) hereby informs you that we will warrant the specified and agreed performance of the Lightsheet Z.1 system only if sample chambers are applied and used that either are delivered or explicitly approved by us.

The sample chamber design has been optimized to ensure the most established applications of Light Sheet Fluorescence Microscopy. Exceptional applications may require a slightly modified sample chamber design. In order to enable customized modifications of the existing sample chamber we also provide the corresponding CAD file and a technical drawing. We explicitly advise you that already minor deviations of the dimensions and tolerances specified in these documents will cause a significant loss of image quality and can potentially result in a liquid leakage. Therefore, you will not hold us or one of our affiliates liable for any damages caused by the employment of self-built or third-party-built sample chambers, the use of such self-built or third-party-built sample chambers will be solely on your own risk. Furthermore we want to inform you, that we will not render any assistance relating to the production and application of such self-built or third-party-built sample chambers.

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CHAPTER 1 - HARDWARE

o cool the

Carl Zeiss User Interfaces Lightsheet Z.1

3.1 Installation and Deinstallation of the Detection Modules

A cooling liquid defined as a hazardous substance is used in the Lightsheet Z.1 t detection modules (depending on configuration). The supplied safety data sheet with notes on hazards and safety measures must be observed when handling the cooling liquid.

The ports of the detection modules, reflection (Cam1) (Fig. 3/1) and transmission (Cam2) (Fig. 3/2), on the main system module Lightsheet Z.1 are equipped with a hardware interlock device. The contact rings (Fig. 3/9) are connected to the detection modules (Fig. 3/8) and adjusted in the factory. When handling the detection modules, ensure that neither tilting nor rotation forces are exerted on this connection to avoid misalignment. The detection module should be gripped as close as possible to the contact ring.

A rotation guard plate (Fig. 3/7) is mounted on the contact ring of each detection module. When mounting on both ports this must point toward the front side of the Lightsheet Z.1. It prevents rotation of the detection module and carries the laser protection notice.

Never remove the rotation guard plates of the detection module adapters. Do not modify or manipulate the detection module adapters.

The lock is functioning when the sensors (Fig. 3/4) of the sensor disk (Fig. 3/11) are pressed down by the pins of the contact ring (Fig. 3/10 or 5 for blind cap). If this is not the case, e.g. the space between the two devices is too large, the laser is blocked and the system cannot be used.

If the system does not function after a detection module has been attached to or removed from a

port with a safety interlock, check the connection of the contact ring on the respective detection module or blind cap (Fig. 3/9 or 6) to the sensor ring.

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1 Detector port 1 (reflection, Cam1)

2 Detector port 2 (transmission, Cam2) 3 Securing screw 4 Sensors (2×) in sensor ring 5 Contact pins (2×) on blind cap 6 Blind cap 7 Rotation guard plate with laser protection notice 8 Detection module (varies) 9 Contact ring of the detection module 10 Contact pins (2×) on contact ring 11 Sensor disk 12 Sensor ring

Fig. 3 Mounting and dismounting the detection modules

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CHAPTER 1 - HARDWARE Carl Zeiss User Interfaces Lightsheet Z.1

The exchange of the detection modules on your Lightsheet Z.1 can be divided into two parts. One deals with the actual mounting and dismounting of the detection modules and their connections to the system. The second part deals with the registration and alignment of the detection module using specific software tools.

The Lightsheet Z.1 must be completely turned off before changing detection modules.

No sample, capillary or similar object must be in the beam path while the alignment and calibration

tasks are performed.

1) Dismounting and Mounting of Detection Modules

− Disconnect all cables from the detection module.

− The connection of the liquid cooling can stay in place, but the power plug for the liquid cooling

unit must be pulled when the detection module is not in use.

− Now disconnect all cables from the Lightsheet Z.1 system and the PC for system control. Even if the detection modules are regularly switched no cable should remain on the system.

− Hold the detection module firmly in one hand.

− Loosen the securing screw with an Allen wrench (Fig. 3/3) on the sensor ring (Fig. 3/12) of the

detector port (Fig. 3/1 and 2).

− Carefully pull away the detection module (Fig. 3/8) with contact ring (Fig. 3/9) using a slight tilting motion if necessary.

− Cover the detector port (Fig. 3/1 and 2) on the main system module Lightsheet Z.1 and on the detection module with the caps provided (Fig. 3/6).

− For mounting the new detection module, proceed in the reverse order. Tighten the securing screw (Fig. 3/3) on the sensor ring (Fig. 3/12) of the detector port (Fig. 3/1 and 2) without applying force.

− With the detection module "PCO.Edge," note that part of the optics projects beyond the contact ring. To protect it from damage and fingerprints, first push the optics into the port and then connect the contact ring to the sensor ring.

− Connect all necessary cables to the detection module, the Lightsheet Z.1 and the PC for system control (see section 3.2.4 Cable Connections for the Detection Module "PCO.Edge" and

3.2.5, Cable Connections for the Detection Module "Standard").

− Use the cable holder (Fig. 3/3) on the system table to connect the cables (Fig. 3/1) of the detection modules without tension.

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Fig.

Lightsheet Z.1 User Interfaces Carl Zeiss

1 Cable (e. g. of the detection modules)

2 Cable tie 3 Cable holder on system table

Fig. 4 Cable strain relief

− Turn on the system (see CHAPTER 4, SYSTEM OPERATION).

2) Registration and Alignment of the detection modules

− After the Lightsheet Z.1 and the PC for system control are turned on and the operation system has

booted, start the software ZEN Configuration Tool

. This can be found either as a shortcut on

your desktop or in this directory: C:\ZEN\HWT as Configuration Tool 2012.exe.

− In the opening window press the Start configuring button (Fig. 5).

5 ZEN Configuration Tool Start-up window

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CHAPTER 1 - HARDWARE Carl Zeiss User Interfaces Lightsheet Z.1

− On the Configuration tab (Fig. 6) you will find a Camera field with a drop – down menu for both detector ports (Cam1, Cam2). Select the detection module(s) you are planning to use. When you use the PCO.Edge, the checkbox liquid cooling will be marked automatically.

Fig. 6 ZEN Configuration Tool, Configuration tab

− Press the Store all Changes button and then exit the Configuration Tool. This will save the changes you have made into the database of your system.

If you plan to exchange detection modules on a regular basis, it can be helpful to create two

databases, one for each detection module type. To do so, copy the original database with a different name (you find it in the directory: C:\ZEN\database) and modify it with the Configuration Tool accordingly. Make sure to always keep the original database. Name both databases, to easily identify them. When ZEN is started open the Boot Status with the black arrow and then the Hardware configuration database the same way (Fig. 7). Here you can choose which database to use. The Recent… button shows all databases that have been used lately. The Choose… button opens a window explorer window which allows one to search for a database to use.

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− Now start the ZEN (black edition) software and perform the following tasks found in the Maintain tab:

• Detector recognition

• Automatic Detector Alignment; and

if necessary

• Manual Detector Alignment.

− It might be necessary to adjust the focus

of the calibration grating (Fig. 8) for the Automatic or Manual Detector Alignment task. Start one of these tasks to get a live image of the grating for evaluating and adjusting the focus if applicable (see section 3.2.2 Adjustment – Manual

Detector Alignment).

Fig. 7 ZEN Start-up window

3.2 Adjustment – Detector Recognition

When the detection modules are exchanged or attached to the Lightsheet Z.1 system for the first time, the system must identify which detection module is present for which channel. Pressing the button Detector recognition will start an identification routine to do this job. A progress bar is visible during the process. When the Cancel button is pressed, the operation is aborted without saving any result. This routine must be conducted before the Automatic or Manual Detector Alignment. At the end of this process you be informed if the detector recognition had to perform any changes. If so, please restart the ZEN software before continuing.

3.2.1 Adjustment – Automatic Detector Alignment

This adjustment is necessary for one or two detection modules, when:

• detection modules have been exchanged

• reflector turrets have been removed and inserted (restart ZEN before the alignment task)

• recognition of a pixel shift between channels

• the zoom is not centered.

No sample should be present. The necessary correction plate and grating is present for each channel within the system and needs no additional activation.

The Automatic Detector Alignment will align the image center with the detection module center (zoom center). For two detection modules, it will additionally align both to have overlapping pixels.

After detection module exchange, first press the Detector recognition button in the Adjustment

tool window.

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After pressing the Automatic Detector Alignment button, a wizard window (Fig. 8) will open that guides you through the process. Press the Next button after each step to continue or the Previous button to move one step back. If you press Cancel you will abort the process without saving any results.

The first step (Fig. 8) automatically positions the necessary grating into the light path and adjusts brightness and contrast. A progress bar is present in the lower part of the window. You can follow the process in the displayed live image. You find the following text below the live image:

“This will start detector alignment for all available detectors and filter positions in the system. Autoadjustment of brightness and contrast, please wait.”

If the live image of the grating shows the

grid pattern out-of-focus, cancel the

Fig. 8 Automatic Detector Alignment, first

step

Automatic Camera Alignment wizard and first manually adjust the focus of the grating. The necessary steps are described in section

3.2.3 Adjust the Grating Focus for the Automatic or Manual Detector Alignment Tool.

In Step 2 the movement of the correction plates is calibrated for both channels. There is no interaction needed. Press the Next button to continue after the task is done. The following text is displayed:

“Calibration of correction plate movement, please wait.”

In Step 3 the zoom center is determined. A graphic of the grating is displayed at the left hand side of the window (Fig. 9). Two squares are labeled with white crosses in this image. On the right hand side, a live image of the grating is shown. Now mark the same squares as shown in the graphic in the live image by simply clicking on them with the left mouse button. This will be repeated for a second zoom step. The following text will be shown: “Determining the detector center. Please click on the two squares in the

right live image of the grid that are shown with the white cross within the left image display. Repeat for both zoom settings, then continue.”

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Step 4 is the final step in which the alignment for all filters and detection module channels is performed. A progress bar is present at the bottom of the window. When the procedure is finished the Finish button becomes available, press it to save all adjustments and leave the wizard. This text is displayed: “Detector Alignment for all available filters in progress, please wait.”

If you still encounter pixel shifts between the two channels after the Automatic Camera Alignment tool, you can fine tune settings using the Manual Detector Alignment tool, see section 3.2.3 Adjust

the Grating Focus for the Automatic or Manual Detector Alignment Tool.

Fig. 9 Automatic Detector Alignment,

third step

3.2.2 Adjustment – Manual Detector Alignment

This adjustment is necessary for one or two detection modules when you recognize a pixel shift between channels, or when the zoom is not centered.

The Automatic Detector Alignment

should always be performed first since it might already solve the problems. Furthermore it defines the settings for the combination Channel 1 (Cam1) and position 1 of the Emission Selection Filter, which should be used as the reference during the Manual Detector Alignment.

The Zoom in the Acquisition tool window

is fixed to 1 while performing the Manual Detector Alignment.

Fig. 10 Manual Detector Alignment

window

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No sample should be present when performing this task. When pressing the Manual Detector Alignment button, a window will open (Fig. 10). A grating is automatically inserted into the light path and a live image of it is shown in a new image container within the ZEN software. The Set Exposure button is activated automatically to adjust brightness. All components of the light path can be controlled in the Manual Detector Alignment window. The light source is the LED white light source to ensure the visibility of the grating independent of the filters in the light path. All other controls for adjusting imaging settings should be controlled in the tool windows of the main software.

The activated (blue) Automatic button loads the settings which have been previously defined with the Automatic Detector Alignment for each light path.

After pressing the Continuous button in the

Manual Detector Alignment window or in the Main tool tabs a new image will open with a live

image of the grating (Fig. 11)

Fig. 11 Manual Detector Alignment,

Grating at Zoom 1

When the grating is out of focus, you need

to refocus it for the relevant channel by performing the steps described in section

3.2.3. To check the focus, zoom into the image using the Zoom function and look at the center cross within the square (Fig. 12). If the white portions within the inner most circle are recognizable, the grating is in focus.

To zoom into the image you can use these buttons:

As a reference, the combination of Emission Selection filter position 1 and Cam 1 should be taken (see above). This combination should be the first live image. Here mark a structure close to the middle of the circled cross using the Graphics View control. This cross will be the reference for the following steps.

Now change the light path to the combination of

Fig. 12 Manual Detector Alignment,

Grating, center part zoomed in

Emission Selection filter and Channel (Cam 1 or Cam 2) that you wish to adjust. The live image will now show the grating with the current settings.

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Use the sliders or the input box with arrows of Cam 1 X and Cam 1 Y (for Channel 1, Cam1) or Cam 2 X and Cam 2 Y (for Channel 2, Cam 2) to move the previously chosen structure of the grating to overlay with the cross. Finish this step by pressing the Store Current Settings button.

Repeat the procedure for all desired Emission Selection filters and channels, always using the cross marker as the reference.

It is helpful, especially after an Automatic Detector Alignment has been performed, to use the Profile View tab in order to evaluate if the image of Cam 1 and Cam 2 overlay. While the grating is continuously imaged, press the Profile View tab and draw a line or an arrow poly-line on the grating (Fig. 13). Use the sliders or the input box with arrows of Cam 1 X and Cam 1 Y (for Channel 1, Cam1) or

Cam 2 X and Cam 2 Y (for Channel 2, Cam 2) to move the lines of the grating to overlay each other.

Fig. 13 Manual Detector Alignment, Grating, Profile view tab

You can leave the Manual Detector Alignment by pressing the Close button at the bottom of the window.

3.2.3 Adjust the Grating Focus for the Automatic or Manual Detector Alignment Tool

During the Automatic Detector Alignment tool wizard and the Manual Detector Alignment tool a grating is brought into the light path which can be imaged on each channel. When the grating is out-offocus, you need to refocus it for the relevant channel. For checking the focus, it helps to zoom into the image using the Zoom function and look at the center cross within the square (Fig. 12) If the white portions within the inner most circle are recognizable, the grating is in focus.

To adjust the focus of the grating start the Manual Detector Alignment by pressing the button in the Adjustment tool window. The grating will be brought into the beam path and an image is generated and displayed in the image container of the main software.

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Set focus for Cam 1 (Channel 1)

• Use the Light Path of the Manual Detector Alignment to first activate channel 1 (Cam1) and press afterwards the Continuous button. The resulting live image shows the grating on channel 1 (Cam1).

• Use the Zoom tools of the Manual Detector Alignment to zoom into the center region of the image.

• With the live image running, open the front system door (Fig. 14/1) to the reflector turret for

emission filter selection. This system door has no safety interlock, and opening it will not terminate image acquisition.

• Turn the screw which is above the turret (Fig. 14/3) with the long Allen wrench (Fig. 14/2).

1 Front system door

2 Allen wrench 3 Screw

Fig. 14 Position of the screw to adjust the focus of the grating for Cam 1 (channel 1)

• Monitor the live image of the grating while you turn the screw in any direction and find the position of the screw that results in the sharpest image.

• Stop the continuous scan and close the front system door.

Set focus for Cam 2 (Channel 2)

• Use the Light Path of the Manual Detector Alignment to direct light towards channel 2 and activate channel 2 (Cam2).

• Press afterwards the Continuous button. The resulting live image shows the grating on channel 2 (Cam2).

• Use the Zoom tools for the Manual Detector Alignment to zoom into the center region of the image.

• While the imaging continues remove the black screw cap (Fig. 15/1) on the back of the main system

module Lightsheet Z.1. This opening gives way to a screw that can be turned with an Allen wrench (Fig. 15/2).

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• Monitor the live image of the grating while you turn the screw in any direction and find the position of the screw that results in the sharpest image.

• Stop the continuous scan and place the black screw cap back on the system.

1 Black screw cap

2 Allen wrench

Fig. 15 Position of the screw to adjust the focus of the grating for Cam 2 (channel 2)

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3.2.4 Cable Connections for the Detection Module "PCO.Edge"

The detection module "PCO.Edge" is equipped with a liquid cooling system. Please read the safety notes of the Lightsheet Z.1 system, as well as the separate safety data sheet for the liquid cooling system. The power supply unit (PSU) for the cooling is connected to the system's power strip at the rear of the laser module. The cooling hoses should be as short as possible. The connection creates a closed circuit, starting from the cooling unit leading to the detection module 1 (approx. 200 cm hose length) reflection port (Cam1). From the latter a hose (approx. 200 cm) leads either back to the cooling unit or, if a second detection module is used, first to the transmission port (Cam2) (approx. 100 cm) and then a third hose (approx. 200 cm) leads back to the cooling unit.

Fig. 16 Cable connections for the detection module "PCO.Edge"

The detection module cables are connected as follows:

− CameraLink cables A and B are each plugged into the PC for system control. The upper slots of the PC for system control are assigned to the detection module of the reflection port (Cam1), [CamLink A] right and [CamLink B] left. The lower slots are assigned to the transmission port (Cam2), [CamLink A[ right and [CamLink B] left.

− The power cable is connected to the power strip of the system on the rear of the LB Rack Lightsheet.

− The trigger cable is always connected to the detection module with [1/IN] and on the rear side of the main system module Lightsheet Z.1 to [Camera 1/IN] (refection port, Cam1) or [Camera 2/IN] (transmission port, Cam2).

− The detection module is switched on with the toggle switch on the module itself.

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3.2.5 Cable Connections for the Detection Module "Standard"

− The following connections exist for this detection module:

− The firewire cable must be plugged into the PC for system control. There are four firewire sockets

on the PC for system control; the detection module from the reflection port (Cam1) is connected to position [FW 1], from the transmission port (Cam2) to position [FW 3].

− The trigger cable is plugged into the respective detection module and the three ends on the rear side of the main system module Lightsheet Z.1 to [Camera 1] or [Camera 2], as well as [Camera 1/IN] and [Camera 1/OUT] (reflection port, Cam1) or [Camera 2/IN] and [Camera 2/OUT] (transmission port, Cam2).

Fig. 17 Cable connections for the detection module "Standard"

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3.3 Assembly of the Sample Chamber

− Wear powder-free gloves to avoid fingerprints on the cover slips as well as to protect your hands from sharp edges.

3.3.1 Assembly of the Sample Chamber Windows

− Assemble the three sample chamber body windows (for the illumination optics on the sides and the front chamber window for the overview camera with LED white illumination) using the following steps:

1 Sample chamber body

2 Cover slip 3 O-ring (18 mm O-ring, black)

4 Illumination adapter ring 5 Sample chamber window tool

Fig. 18 Assembly of the three sample chamber windows for illumination optics and overview

camera

1. Take the sample chamber body (Fig. 18/1) and lay it onto the table so that one of the three smaller windows are facing you.

2. Using a forceps or gloved hands, remove one clean circular 18 mm cover slip; place cover slip into the empty sample chamber body window so it fits into the smallest groove (Fig. 18/2) – you will need a fine forceps.

3. Now place the O-ring (18 mm O-ring, black) into the corresponding groove (Fig. 18/3). Make sure not to disturb the positioning of the cover slip.

4. Take the silver illumination adapter ring (Fig. 18/4) and lay it into the window making sure the flat side with the four pinholes is facing you.

5. Using the sample chamber window tool (Fig. 18/5), position the circular pins into the pinholes and turn clockwise until finger-tight. Adapter should be flush with the sample chamber body.

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6. Repeat step 1 - 6 for each of the three windows.

− Assemble the sample chamber window for the detection optic.

1. Turn the sample chamber body (Fig. 19/1) to the largest opening window for the detection optics.

2. Using a small spatula or the opposite end of a forceps position the blue O-ring tightly into the groove to ensure no leaking (Fig. 19/2).

3. Assemble the chamber window for the detection optic according to the following steps. Depending on the detection optics used, insert the correct detection optic adapter:

4. For the detection optic 5x/0.16:

a) Place the large silver detection optic adapter 5x (Fig. 19/3) onto the table with the

grooves facing you.

b) Remove one clean circular 18 mm glass cover slip; place cover slip (Fig. 19/4) into

the large silver detection optic adapter 5x (Fig. 19/3) making sure it is positioned into the smallest groove – adjust the coverslip position with the forceps if necessary.

c) Now place the O-ring (18 mm O-ring, black) into the corresponding groove on top

of the cover slip (Fig. 19/5).

d) Take the silver retaining ring (Fig. 19/6) and place it inside the detection optics

adapter 5x with the flat side with two notches facing you. Using the circular pin-side of the sample chamber window tool (Fig. 19/9), turn clockwise until finger-tight.

e) Placing the assembled detection optic adapter 5x into the sample chamber body

window. Using the squared pins of the sample chamber window tool, turn clockwise until finger-tight.

5. For the detection optic 20x/1.0, 40x/1.0, 63x/1.0:

a) Use the corresponding detection optic adapter (see the detection optic adapter

catalogue number and magnification engraving) (Fig. 19/7).

b) Insert the detection optic adapter into the sample chamber body window for

detection with the flat side facing you.

c) Using the sample chamber window tool (squared pin-side) (Fig. 19/9) turn adapter

clock-wise until finger-tight.

d) Take a 15 mm black O-ring (Fig. 19/8) and position the O-ring into the groove inside

the middle of the detection optic adapter using the blunt end of a forceps if necessary.

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1 Sample chamber body

2 O-ring (blue) 3-6 For the detection optic 5x/0.16

− 3 Detection optic adapter 5x/0.16

− 4 Cover slip

− 5 O-ring (18mm O-ring, black)

− 6 Retaining ring 7-8 For the detection optic 20x/1.0, 40x/1.0, 63x/1.0

− 7 Detection optic adapter (20x/1.0, 40x/1.0, 63x/1.0

− 8 O-ring (15mm O-ring, black) 9 Sample chamber window tool

Fig. 19 Assembly of sample chamber window for the detection optic

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3.3.2 Assembly of the Sample Chamber Body and the Sample Chamber Dove Tail Slide

1 Mesh screen 2 Sample chamber body

Fig. 20 Placing the mesh screen

− Place the mesh screen (Fig. 20/2) into the middle of the sample chamber body (Fig. 20/1) to reduce turbulence stress on the specimen when the chamber is filled.

− For cleaning, it can be carefully removed by a forceps or via a teflon-tip-free end of a capillary plunger.

− If a Heating component is used, proceed first with chapter 3.7.1 (see PeCon manual for specific heating component) and screw it onto the bottom of sample chamber body.

• The sample chamber body (Fig. 21/3) can now be attached to the sample chamber dove tail slide (Fig. 21/1).

• Lay down the sample chamber dove tail slide with the two positioning pins facing upwards (Fig. 21/2).

• Take the sample chamber body and position it on to the sample chamber dove tail slide with the

detection optic adapter window facing to the back relative to the sample chamber dove tail slide grip.

• Invert the sample chamber body with the sample chamber dove tail slide so that the four screw-holes (Fig. 21/4) that were on the bottom are now facing you.

• Insert the four screws (diameter = M3, length = 14 mm) (Fig. 21/5) into the four screw-holes and tighten using and Allen wrench (2.5).

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1 Sample chamber dove tail slide 2 Positioning pins 3 Sample chamber body 4 Screw-holes 5 Screws (diameter = M3, length = 14 mm)

Fig. 21 Assembly of the sample chamber body and the sample chamber dove tail slide

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3.3.3 Insertion of the Drain Connector, Luer-Lock Connectors and Blind Plugs

− Turn the assembled sample chamber body with dove tail slide so that the dove tail slide is on the

table with the grip facing you (Fig. 22/1).

− Take the grey drain connector and screw this into the upper left hole of the sample chamber body

until finger-tight (Fig. 22/6). This will guide excess liquid to the drain.

− Take a Luer-Lock-connector (white) and screw it into the lower right corner of the sample chamber

body (Fig. 22/4). The Luer Lock connectors (white) are used to fill or perfuse the sample chamber with liquids and / or gasses.

− For CO

incubation, the Luer-Lock connector must be positioned at the upper right corner of the

sample chamber (Fig. 22/2). For syringe based perfusion one can use any of the three lower screw holes. The lower right corner screw hole is recommended for convenience (Fig. 22/4).

− For perfusion pump use the right side bottom screw hole for the perfusion input and either of the

front lower corner screw holes for the perfusion output.

− Close all remaining sample chamber screw hole openings by inserting the black blind plugs

(Fig. 22/3; Fig. 22/5).

1 Sample chamber body with dove tail slide 2 Luer-Lock-connector (white); position recommend for CO 3 Blind plug (black) 4 Luer-Lock-connector (white); position recommend for syringe based perfusion 5 Blind plug (black) 6 Drain connector (grey)

Fig. 22 Insertion of the Drain connector, Luer-Lock-connectors and Blind plugs

incubation

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3.3.4 Insertion of Accessories for Incubation

• A temperature sensor (Fig. 23/1) can be used in the sample chamber body.

− Hold the temperature sensor so that it shows you the letter “J”. Slide the temperature sensor

sleeve (Fig. 23/3) onto the top horizontal part of the J-shaped temperature sensor.

− Dip the temperature sensor with sleeve into the sample chamber body and position the temperature sensor sleeve securely into the sample chamber body groove (Fig. 23/2).

1 Temperature sensor 2 Sample chamber body groove 3 Temperature sensor sleeve

Fig. 23 Insertion of temperature sensor

− Place a cover on top of the sample chamber body (Fig. 24/3). This will minimize evaporation of medium or contamination of the sample chamber. If CO

module Lightsheet Z.1 is used

(configuration dependent, chapter 2.7.2) the cover is recommended to maintain the CO concentrations.

− Keep the opening as small as possible. Use the 3 mm opening cover for capillaries (Fig. 24/1) and

7 mm opening cover for syringes (Fig. 24/2). The cover only rests on top of the sample chamber body and is free to move.

gas

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1 Cover for capillaries (3 mm opening) 2 Cover for syringes (7 mm opening) 3 Sample chamber body with dove tail slide

Fig. 24 Placing cover on top of the sample chamber body

3.4 Removing and Inserting the Sample Chamber

The sample chamber can be removed from the system. For removal, proceed as follows:

− Open the front system cavity door (Fig. 25/1) of the main system module Lightsheet Z.1.

− Remove the liquid from the sample chamber (Fig. 25/6) using the hose with the corresponding

syringe (Fig. 25/9).

− Ensure that no sample protrudes into the sample chamber. Remove the sample, if necessary, through the upper sample opening (Fig. 25/2) below the upper system cavity door (Fig. 25/3).

− If necessary, remove the connections for the incubation device (Fig. 25/8).

− Loosen the screw (Fig. 25/4) on the sample chamber and pull the sample chamber out by the grip

(Fig. 25/5).

− Inserting the sample chamber:

− Take hold of the sample chamber by the lower grip (Fig. 25/5) and push the sample chamber

(Fig. 25/6) into the guide rails (Fig. 25/7).

− Tighten the screw (Fig. 25/4) without exerting force. The sample chamber has been correctly inserted when its lower edge is flush with the guide rail.

− Plug in the cable to the incubation unit (Fig. 25/8) as necessary.

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1 Front system cavity door 6 Sample chamber 2 Upper sample opening 7 Guide rails 3 Upper system cavity door 8 Connections for incubation 4 Securing screw 9 Hose and syringe 5 Sample chamber grip

Fig. 25 Removing and inserting the sample chamber

/ sample chamber mount

3.5 Assembly of the Sample Holder

Depending on the size of the sample there are two different types of sample holders available: sample holder for capillaries (inner diameter of capillary size 1 / ~0.68 mm, size 2 / ~1 mm, size 3/ ~1.5 mm, size 4 / ~2.15 mm) and syringes (1 ml). Always use the minimal cylinder diameter necessary for your specimen size to avoid excessive amounts of embedding medium.

3.5.1 Assembly of the Sample Holder for Capillary

1. Select the corresponding colored sleeves (Fig. 26/3) to match the color of the capillary of choice (Fig. 26/6).

2. The capillary should hold the sample and the appropriate plunger (Fig. 26/5), see as well the CHAPTER 2 SAMPLE PREPARATION in this manual. The plungers that fit into capillary size 2 - 4 have to be used with corresponding Teflon tips that are already assembled onto the plungers. Note that in addition 10x Teflon tips of each as well as matching Teflon tip tools are provided in the Chamber & Sample Holder Starter Kit Lightsheet Z.1 (Tab. 1). The Teflon tips can be assembled as a replacement onto the matching tip-less plungers.

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Capillary size (inner diameter)

Capillary Color coding/

Plunger Reference number

Teflon tip Teflon tip too

Color coding

Reference number

Size 1 (~0.68 mm) Red/701902 #701996 - -

Size 2 (~1mm) Black/701904 #701997 Size 2 Clear

Size 3 (~1.5mm) Green/701908 #701998 Size 3 Green

Size 4 (~2.15mm) Blue/701910 #701999 Size 4 Blue

Tab. 1 Capillary components in the Chamber & Sample Holder Starter Kit Lightsheet Z.1

3. The sleeves are tube-shaped with four slits at one end. Insert the sleeves into the capillary sample holder stem (Fig. 26/2) so that the two slit endings point outwards and the non-slit endings face toward the center.

4. Take the clamp screw (Fig. 26/4) and position it onto the capillary sample stem holder turning clockwise three times (360° each turn).

1 Sample holder disc for capillaries 5 Plunger 2 Capillary sample holder stem 6 Capillary with color-coded marking 3 Sleeves (color-coded) 7 Ejection tool 4 Clamp screw

Fig. 26 Assembly of the sample holder for capillary

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5. Take the capillary by the glass portion (avoiding the plunger) with the agarose embedded specimen inside and carefully place it into the center of the clamp screw sample holder top.

6. Push the capillary within the holder downwards until the color-coded capillary marking becomes visible (Fig. 26/6 right hand side).

Newly bought sleeves can be slightly tight at their slit opening ends so that in can be difficult to

push the capillary within the holder. In this case take the capillary with matching plunger and insert it carefully into the corresponding sleeves at their non-slit endings. Make sure that the slits point outwards. Position the capillary surrounded by the sleeves into the sample holder stem and place the clamp screw. You can proceed with step 6).

7. Tighten the clamp screw. Take the sample holder disc (Fig. 26/1) with the protruding side with positioning notch facing you. Carefully position the capillary stem holder with specimen into the sample disc holder until the ball bearing click-position is felt.

8. Before taking the capillary out (chapter 2.6), retract the sample back inside the capillary glass via using plunger.

9. For removal, loosen the clamp screw and carefully pull the capillary through the stem sleeve by using the glass end nearest the color-coded marking.

10. Unscrew the clamp screw completely and remove the sleeves by using the ejection tool (Fig. 26/7) for the capillary sleeves removal.

The sleeves as well as the glass portion of the capillary can be cleaned with a lint-free cloth moistened with water.

3.5.2 Assembly of Sample Holder for Syringes

− Place the sample holder disc for syringes (Fig. 27/1) so the flat side is facing you.

− Take the adapter ring (Fig. 27/2) and place it into the center of the sample holder disc with the

larger diameter of the ring facing you.

− Pick up the sample disc holder with adapter ring in one hand. Take the syringe carrying your specimen (Fig. 27/3) and insert it completely into the sample disc holder (avoiding touching the plunger).

− Touching the syringe body and turn clock-wise to position the syringe body into the two clamps (Fig. 27/6).

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1 Sample holder disc for syringes 4 Ball bearings 2 Adapter ring 5 Syringe body 3 Syringe 6 Clamp

Fig. 27 Assembly of sample holder for syringes

3.6 Inserting and Removing the Sample Holder

• To insert the sample holder move the stage of

the Lightsheet Z.1 system to the Load position via the Specimen Navigator tool (Fig. 28) in the ZEN software (see CHAPTER 4 SYSTEM OPERATION).

• Open the upper system cavity door (Fig. 28/1).

• Take the sample holder for capillaries or syringes by the sample holder disc with the capillary or the

syringe facing downwards (Fig. 29/3).

Fig. 28 Load Position in the Specimen

Navigator

• Insert the sample holder gliding the sample holder disc along the guide rails (Fig. 29/2). It is placed

correctly into a click position if the sample holder disc ball bearings lock into the three holes.

• Notice the white line marking at the outer edge of the disc. It can be used as a reference point for

reorienting the sample holder disc once it was taken out.

• Close the upper system cavity door (Fig. 29/1) and position the capillary and the sample as described in

CHAPTER 4 SYSTEM OPERATION.

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1 Upper system cavity door 2 Guide rails 3 Capillary 4 Hole for disc ball bearings

Fig. 29 Inserting the Sample Holder

− Before taking the sample holder out, retract the sample back inside the capillary glass via using plunger.

− Move to the Load position via the Specimen Navigator tool (Fig. 28) in the ZEN software (see CHAPTER 4 SYSTEM OPERATION).

− Open the upper system cavity door (Fig. 29/1).

− Take the sample holder out by gliding the sample holder disc along the guide rails (Fig. 30).

Fig. 30 Removing the Sample Holder

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3.7 Installation of the Incubation Modules

The incubation of the Lightsheet Z.1 is configuration dependent. It is used to control and monitor temperature and CO various configurations.

− The configuration dependent modules should be placed on a level surface next to the system and are stacked on top of each other (Fig. 31). For further information see the individual component manuals from PeCon.

The installation of incubation modules on the Lightsheet Z.1 can be divided into two parts. One deals with the actual assembly of hardware components of the incubation modules and their connections to the system. The second part deals with the registration of the incubation modules using specific software tools.

supply to the sample environment. The TempModule S1 is required for all of the

Fig. 31 Incubation modules stacked on top

of each other

3.7.1 Heating Components

There are two different heating components available: (1) Heating Block Sample Chamber Lightsheet Z.1 enables to heat the sample chamber from ambient temperature to 42 °C. (2) Peltierblock Sample Chamber Lightsheet Z.1 permits heating and cooling of the sample chamber in the range from 10 °C to 42 °C.

(1) Heatingblock Sample Chamber Lightsheet Z.1 is controlled by the TempModule S1 providing electrical power supply and closed-loop temperature control. The system allows to heat the sample chamber from ambient temperature to 42 °C.

Connect the Heating Block LSFM and the TempModule S via connecting cable. For proper electrical

connection please follow the connection diagram in the individual component manuals from PeCon.

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Make sure that the temperature inside the sample chamber is in accordance with the desired

temperature value required for the experiment. The ZEN software displays the temperature of the heating block itself and does not register the actual temperature in the sample chamber that can be lower (due thermal loss) or higher.

• To assemble the heating block to the sample chamber remove first of all the sample chamber dove tail slide from the assembled sample chamber (Fig. 1, section 3.3.2 Assembly of the Sample Chamber Body and the Sample Chamber Dove Tail Slide).

• Take the heating block and screw it onto the bottom of sample chamber body following the instructions of the individual component manuals from PeCon.

• The sample chamber body can now be assembled together with the sample chamber dove tail slide (Fig. 1, section 3.3.2).

• Insert the sample chamber as described in chapter 2.4 and plug in the 8 pin connector of the connection cable into the right side of the front system cavity / sample room following the individual component manuals from PeCon (Fig. 32/6) shows the connection exemplary for the Peltier Block S).

(2) Peltier Block S is part of the TempModule CZ-LSFM. The Peltier Block helps adjust the sample chamber temperature by heating or cooling the sample chamber within the range of 10 °C to 42 °C (up to 1.5°/min heating, up to 1.0°/min cooling). The temperature inside the sample chamber is monitored by a temperature sensor (Fig. 23/1; Fig. 32/3). For insertion of the temperature sensor in the sample chamber refer to section 3.3.4 Insertion of Accessories for Incubation).

The TempModule CZ-LSFM contains a water based coolant reservoir. For safety reasons (Risk of

electrical shock as a result of leakage) the unit must be placed directly on a level table and is thus the base for all other incubation related modules (Fig. 31).

− Connect the TempModule S, the TempModule CZ-LSFM, the Lightsheet Z.1 and the Peltier Block

according to the individual component manuals from PeCon.

− To assemble the Peltier Block to the sample chamber remove first of all the sample chamber dove

tail slide from the assembled sample chamber (section 3.3.2; Fig. 21).

− Take the Peltier Block and screw it onto the bottom of sample chamber body following the

instructions of the individual component manuals from PeCon.

− The sample chamber body can now be assembled together with the sample chamber dove tail slide

(Fig. 21) and insert it into the Lightsheet Z.1 as described (section 3.3.2 ).

− Insert the accessories for incubation (temperature sensor, cover recommended) into the sample

chamber (section 3.3.4, Fig. 23, Fig. 24).

− Connect the Peltier Block (Fig. 32/6) to the Lightsheet Z.1 by the two provided silicone tubes

(Fig. 32/7-8) and plug in the 8 pin connector of the connection cable into the right side of the front system cavity / sample room (Fig. 32). For proper electrical connections and tube connections follow the instructions of the PeCon manual.

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Heating component Temperature range Incubation modules

Heating Block Heating from ambient

temperature to 42 °C

Peltier Block Heating and cooling in the

range from 10 °C to 42 °C

(1) TempModule S1

(2) Heating Block LSFM

(1) TempModule S1

(2) TempModule CZ-LSFM (including Peltier Block and Temperature sensor)

Tabelle 1 Overview Heating components

3.7.2 CO

The CO

-Module Lightsheet Z.1 controls and monitors adjustable CO2 concentration (from 0 % to 10 %)

in the sample chamber and requires CO

-Module Lightsheet Z.1

supply, sterile filter and Humidifier S1 to provide CO2-enriched

air.

− At least 3h (or more) may be needed until a stable pH value is reached due to the small surface area in relation to sample chamber volume.

− For assembly of hardware components of the CO

Module and its connections to the system, as

well as the calibration procedure needed follow the instructions from the PeCon manual.

− After CO

calibration, the Lightsheet Z.1 system must be completely turned off before work with

the system can continue.

− Position the Luer-Lock connector for CO

incubation at the top right corner of the sample chamber.

Place a cover on top of the sample chamber body (Fig. 24, chapter 3.3.4) to maintain the CO concentrations.

− The sterility of the CO sample chamber by following the cleaning instruction of all parts of the CO

-enriched air is given by the sterile filter. Take control of the sterility in your

-module (see PeCon

manual) as well as the components of the sample chamber (see CHAPTER 1 HARDWARE).

Please clamp all incoming or outgoing tubing (Fig. 32/2, Fig. 32/5, Fig. 32/9, Fig. 32/10) into the

provided clamping grooves. This will avoid possible disconnection problems after closing the front system cavity door.

Note that for the sake of completeness Fig. 26 shows the connection diagram for the usage of a

perfusion pump (Fig. 32/5, Fig. 32/9). Alternatively a syringe based-perfusion can be used (position Fig. 32/5). Consequently, Fig. 32/9 has to be replaced by a bling plug.

gas

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1 Drain connector 2 Luer-Lock connector for CO 3 Temperature sensor with 4-pin connection cable 4 Blind plug 5 Luer-Lock connector for perfusion pump-in with tubing (or alternatively for syringe based perfusion) 6 Peltier Block S with 8-pin connector 7 Cooling liquid (Silicon tube with male connector) 8 Cooling liquid (Silicon tube with male connector) 9 Luer-Lock connector for perfusion pump-out with tubing 10 Drain connector with tubing 11 Drain tray

incubation with tubing

Fig. 32 Incubation connections for sample chamber body

-Module Lightsheet Z.1 can be extended with Heating Device Humidity S1 to improve thermal

equilibrium in the sample chamber.

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3.7.3 Heating Device Humidity S1

For a better humidification of incubation atmosphere the Humidifier of the corresponding CO

-Module

Lightsheet Z.1 can be inserted into the Heating Device Humidity S1. The Heating Device Humidity S1

enables to improve thermal equilibrium in the sample chamber and avoid condensation in the tubing.

3.7.4 Registration of the incubation modules

• After the Lightsheet Z.1 and the PC for system control are turned on and the operation system has booted start the software ZEN Configuration Tool. You find it either as a shortcut on your desktop or in this directory : C:\ZEN\HWT as Configuration Tool 2012.exe.

• On the opening window press the Start configuring button (Fig. 33):

Fig. 33 ZEN Configuration Tool Start-up window

• On the Configuration tab (Fig. 34) you find an Incubation Components field with check boxes for

Module and TempModule that you can activate or deactivate depending on your incubation

configuration. These modules will show up later as Atmosphere (%) panel and Temperature (

panel respectively in the ZEN software (see CHAPTER 4 SYSTEM OPERATION).

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Fig. 34 ZEN Configuration Tool Incubation Components field

• A drop-down menu enables to assign heating components (Heatingblock, Peltier Block) as well as the Heating Device Humidity to four distinct channels.

Make sure that the assignment of incubation components to the corresponding channels in the

Configuration Tool is in accordance with the electrical connection of the incubation components into the TempModule S1 (“Channel 1-4”, “Control Sensor”).

• We recommend to assign Channel 1 to the heating component (Heatingblock, Peltier Block) and Channel 4 to the Heating Device Humidity if available.

• Do not check the check boxes for external Sensor as well as an Humidity Sensor. These components are not part of the Lightsheet Z.1 incubation modules.

• Leave the Configuration Tool with the Store all Changes button. This will write all changes into the database of your system.

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Lightsheet Z.1 illumination optics 5× / 0.1

Lightsheet Z.1 illumination optics 10× / 0.2

Lightsheet Z.1 User Interfaces Carl Zeiss

3.8 Switch Incubation ON and OFF

• Before you turn ON the incubation, all incubation components you wish to use during the imaging session must be attached to the Lightsheet Z.1 system before starting ZEN software.

• Set the "Incubation" switch on the remote control to the ON position. Make sure that all incubation components are switched on.

• Once ZEN is running you can detach and reattach components.

• To turn OFF the incubation, uncheck the corresponding checkboxes in the Incubator tool window

(see CHAPTER 4 SYSTEM OPERATIONl) or deactivate the incubation components by setting the "Incubation" switch to the OFF position.

Be aware that all incubation components will continue working with the in Incubator tool window

specified parameters after ZEN has been shut down or stopped working. This will keep your sample alive under precisely defined conditions, even if ZEN stops unexpected during an experiment.

3.9 Illumination and Detection Optics

In the Lightsheet Z.1, one or a pair of illumination optics plus one detection optic are always required.

We recommend the following combinations:

Lightsheet Z.1 detection optics 5× / 0.16

Lightsheet Z.1 detection optics 20× / 1.0

Lightsheet Z.1 detection optics 40× / 1.0

Lightsheet Z.1 detection optics 63× / 1.0

yes no

no yes

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Detection Optic

0.36x

0.7x

1.0x

2.5x

Detection Optic

0.36x

0.7x

1.0x

2.5x

Carl Zeiss User Interfaces Lightsheet Z.1

The area that can be imaged with the combination of detection optics with the detection module on the Lightsheet Z.1 is summarized *

in the following tables:

Zoom

x (µm) y (y

*2)

µm) x (µm) y (µm) x (µm) y (µm) x (µm) y (µm)

5× / 0.16 4815 4815 2496 2496 1733 1733 693 693

20× / 1.0 1204 1204 624 624 433 433 173 173

40× / 1.0 602 602 312 312 217 217 87 87

63× / 1.0 382 382 198 198 138 138 55 55

Tabelle 2 Detection module PCO.Edge

Zoom

x (µm) y

*2)

(µm) x (µm) y

*2)

(µm) x (µm) y (µm) x (µm) y (µm)

5× / 0.16 4157 5570 2138 2864 1497 2005 599 802

20× / 1.0 1039 1392 534 716 374 501 150 201

40× / 1.0 520 696 267 358 187 251 75 100

63× / 1.0 330 442 170 227 119 159 48 64

Tabelle 3 Detection module Standard

*1) Typical values for Lightsheet Z.1 systems

*2) Be aware, a homogenous light sheet illumination with full intensity is limited to about 2800 µm in the y direction. Above

and below this center, the illumination intensity will fade towards the edges in the y direction.

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3.9.1 Removing and Inserting the Illumination Optics Unit

The illumination optics unit (Fig. 35/2) can easily be removed and inserted after the removal of the sample chamber (see section 3.4 Removing and Inserting the Sample Chamber).

To protect the front lenses, secure the appropriate protective caps (Fig. 35/3) over each of the illumination optics (Fig. 35/2) and protective cap (Fig. 35/4) over the detection optic (Fig. 35/5). The illumination optics can then be unscrewed by turning anticlockwise.

To insert the illumination optics, proceed in the reverse order by screwing it clockwise into the unit until finger-tight.

Finally, the protective caps can be removed and the sample chamber inserted.

After the illumination optic has been exchanged, the currently used optic must be registered in the ZEN software. The necessary software interface can be found in the Objectives tool window under the Maintain tab.

1 Front system cavity door 4 Protective cap for detection optics unit

2 Illumination optics (2×) 5 Detection optics unit 3 Protective cap for illumination optics unit (2×) 6 Threaded shaft for illumination optics unit (r and l)

Fig. 35 Removing and inserting the illumination optics

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3.9.2 Removing and Inserting the Detection Optics Unit

In order to remove the detection optics unit (Fig. 36/3) located in the rear zone of the system cavity, the sample chamber must first be removed (see Section 3.4). To prevent damage to the illumination optics units, these must also be removed (see Section 3.9.1). Secure the appropriate protective cap (Fig. 36/4) onto the detection optics unit.

Grab the detection optics unit as close as possible to the base and remove it by turning anticlockwise. Caution: the detection optics unit is heavy.

To insert the detection optics unit, it must be screwed in a clockwise direction without exerting force. Subsequently the protective cap should be removed before re-inserting the illumination optics unit and the sample chamber.

After the detection optic has been exchanged, the currently used optic must be registered in the ZEN software. The necessary software interface can be found in the Objectives tool window under the Maintain tab.

Each detection optic unit is labeled with an individual serial number. It ensures that the specific calibration file for this lens, necessary for optimal performance, is used when identified within the software. The calibration file is created by a Carl Zeiss service personnel representative during system installation, or when a detection optic is newly purchased. Ensure that the correct detection optic unit is chosen in the ZEN software interface, based on its name and the specific serial number of the detection optic in use.

1 Front system cavity door

2 Threaded shaft for detection optics unit 3 Detection optics unit 4 Protective cap

Fig. 36 Removing and inserting the detection optics

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3.10 Removing and Inserting the Reflector Turret for Emission Selection

The reflector turret (Fig. 37/5) can be accessed by opening the left-hand-side front system door (Fig. 37/1).

After the reflector turret for emission selection has been exchanged, the Automatic Detector

Alignment should be performed. The ZEN software should be restarted if the reflector turret was

removed, while ZEN was still running.

To remove, first of all turn the silver-colored lever (Fig. 37/2), then pull-out the reflector turret by using the grip (Fig. 37/6).

When inserting, it is recommended that the reflector turret first of all be held by the short grip (Fig. 37/5) and inserted into the guide rail (Fig. 37/3) and then pushed in as far as it will go using the long pull rod (Fig. 37/6). Return the locking lever to its original position. Finally, close the system door.

1 Front system door 4 Reflector turret

2 Silver-colored lever 5 Short grip 3 Guide rail 6 Grip

Fig. 37 Removing and inserting the reflector turret for emission selection

The filters are not push and click filters and cannot be taken out as a whole. The filter cube and the

beam splitter were installed and aligned in the factory. Do not try to take out single filter cubes or beam splitters, it will damage the filters and the alignment, rendering the system in the worst case unusable.

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Fig.

Carl Zeiss User Interfaces Lightsheet Z.1

The emission filters can be exchanged on the filter turret (Fig. 37/4).

− Use the provided tool to loosen the retaining ring around the emission filter you wish to exchange and remove the ring.

− Take out the emission filter, best when wearing powder-free laboratory gloves to prevent fingerprints on the filter, by carefully tilting the filter wheel.

− Insert the new emission filter, the arrow mark on its edge must indicate the direction of the light beam. If no arrow marking is present, the reflective side of the filter must point towards the light source.

− Put the ring on top of the filter and tighten it with the same tool without applying force.

The emission filters facing towards the side of the turret are part of the channel 1 (Cam1, Emission Filter 1) light path; the filters facing the top are part of channel 2 (Cam2, Emission Filter 2).

After replacing the reflector turret for emission selection with new filters, the filter modules must be registered into the database using the Configuration Tool software (Fig. 38) or modified in the ZEN software under the Maintain tab, Filters tool window. The position number on the turret is on the left side of the filter. These numbers correspond with the numbering in the ZEN software.

38 Configuration Tool, Filters tab

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If you plan to exchange the reflector turret

for emission selection on a regular basis, it can be helpful to create two databases, one for each reflector turret. To do so, copy the original database with a different name (you find it in the directory: C:\ZEN\database) and modify it with the Configuration Tool accordingly. Name both databases, to easily identify them. When ZEN is started open the Boot Status with the black arrow and then the Hardware configuration database the same way (Fig. 7). Here you can choose which database to use. The Recent… button shows all databases that have been used lately. The Choose… button opens a window explorer window which allows one to search for a database to use.

Fig. 39 ZEN Software, Maintain tab, Filters

tool window

3.11 Removing and Inserting the Reflector Turret for Laser Blocking Filter

To remove the reflector turret for laser blocking filter (Fig. 40/7), first open the upper rear system door (Fig. 40/4). Beneath this is an additional cover plate (Fig. 40/3) on which the knurled screws left and right (Fig. 40/1) can be loosened manually to lift off the cover by the black grip (Fig. 40/2).

Turn the silver-colored lever (Fig. 40/5) and pull the grip (Fig. 40/9 and 8) upwards to release the reflector turret.

To insert the reflector turret, slide it down using the guide rail (Fig. 40/6) as far as it will go and return the silver-colored lever to its original position. Place the cover over the opening and tighten the knurled screws carefully but without force. Close the upper rear system door.

When the knurled screws are opened, the laser beam will be deactivated by the safety control. In

the event that no laser beam appears after replacing the filter wheel, check the knurled screws.

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You can remove and replace filter modules by push and click.

How to uninstall a module:

− Disengage the filter module from the upper spring elements of the filter insert by tilting it forward, then lift it off the lower spring elements and remove the module.

How to install a module:

− Insert the filter module with the aid of its mounting elements on its left and right into the lower spring clips on the filter insert.

− Press the filter module against the upper spring clips until it engages firmly.

1 Knurled screws (2×) 6 Guiding rail 2 Cover plate grip 7 Reflector turret 3 Cover plate 8 Short grip 4 Upper (rear) system door 9 Grip 5 Silver-colored lever

Fig. 40 Removing and inserting the reflector turret for laser blocking filter

After replacing the reflector turret for laser blocking filter or individual filter cubes, it must be registered in the database with the Configuration Tool software or modified in the ZEN software under the Maintain tab within the Filters tool window (Fig. 42). The position number on the turret is on the left side of the filter. These numbers correspond with the numbering in the ZEN software.

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Fig.

Lightsheet Z.1 User Interfaces Carl Zeiss

41 Configuration Tool, Filters tab

42 ZEN software, Maintain tab, Filters

tool window

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4 Index

Automatic Detector Alignment ...................... 15

Capillary color coding .................................... 33

Capillary size ................................................. 33

Customized Sample Chamber .......................... 9

Detection Modules

Adjust Focus of Grating ............................. 19

Alignment ................................................. 13

Cable Connections "PCO.Edge" ................ 22

Cable Connections "Standard" .................. 23

Deinstallation............................................. 10

Dismounting .............................................. 12

Installation ................................................. 10

Mounting .................................................. 12

Registration ............................................... 13

Detection optics ............................................ 43

Exchange ................................................... 46

Serial number ............................................ 46

Detector Alignment ....................................... 15

Detector Recognition ..................................... 15

ErgoDrive Operating Panel ........................... 6, 7

Illumination optics

Exchange ................................................... 45

Illumination Optics ........................................ 43

Image

Pixel shift ................................................... 15

Zoom not centered .................................... 15

Image Area ................................................... 44

Incubation

-Module .............................................. 39

Cover ........................................................ 30

Heating Components ................................. 37

Heating Device Humidity S1 ....................... 41

Heatingblock ............................................. 37

Installation of Incubation Modules .............. 37

Peltier Block S ............................................ 38

Registration of modules ............................. 41

Temperature sensor ................................... 30

Tempmodule CZ-LSFM ............................... 38

Manual Detector Alignment ...........................17

Plunger reference number ..............................33

Reflector Turret for Emission Selection

Database configuration ...............................49

Exchange ...................................................47

Reflector Turret for Laser Blocking Filter

Exchange ...................................................49

Push and click filter exchange .....................50

Rotate around Center of image ....................... 7

Sample Chamber

Assembly....................................................24

Assembly of Sample Chamber Body ............27

Assembly of Sample Chamber Dove Tail Slide

..............................................................27

Assembly of Sample Chamber Windows .....24

Blind Plugs .................................................29

incubation connection .........................29

Cover .........................................................30

Customized ................................................. 9

Detection optic window ..............................25

Drain Connector .........................................29

Incubation Accessories ................................30

Inserting .....................................................31

Luer-Lock Connectors .................................29

Removing

................................

...................31

Syringe based perfusion ..............................29

Temperature sensor ....................................30

Sample Holder

Assembly....................................................32

Capillary .....................................................32

Inserting in System .....................................35

Plunger ......................................................32

Removing from System ...............................35

Sleeves for Capillary ....................................32

Syringe .......................................................34

Specimen Navigator tool ................................. 7

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CHAPTER 2 - SAMPLE PREPARATION Lightsheet Z.1 Content Carl Zeiss

CHAPTER 2 SAMPLE PREPARATION

Flood P.M., Kelly R., Gutiérrez-Heredia L. and E.G. Reynaud

School of Biology and Environmental Science, University College Dublin, Belfield, Dublin 4, Dublin, Ireland

CONTENT

Page

1 INTRODUCTION ................................................................................................... 3

2 SAMPLE MOUNTING FOR LSFM .......................................................................... 6

2.1 The perfect Sample for LSFM ........................................................................................ 6

2.2 Holding the Sample ....................................................................................................... 8

2.2.1 Embedded Samples ........................................................................................................ 10

2.2.2 Hanging Samples ........................................................................................................... 14

2.2.3 Enclosed Samples ........................................................................................................... 15

2.2.4 FEP Tubing ..................................................................................................................... 17

2.3 Materials and Equipment ............................................................................................ 18

2.3.1 Sample Chambers .......................................................................................................... 18

2.3.2 Molding and Mounting Supports .................................................................................... 20

2.3.3 Sample Holder ............................................................................................................... 22

2.3.4 Gels and Polymers .......................................................................................................... 23

2.3.5 Hydrogel Preparation ...................................................................................................... 24

2.4 Fixation and Fixatives ................................................................................................. 25

2.5 Stains and Staining ..................................................................................................... 25

2.5.1 Choosing a Fluorescent Label ......................................................................................... 26

2.6 Antifading Agents ....................................................................................................... 26

2.7 Cleaning, Labelling and Storing Samples ................................................................... 26

3 SPECIFIC EXAMPLES OF SAMPLE PREPARATION ............................................. 28

3.1 Preparation of Fluorescent Beads ............................................................................... 28

3.2 Preparation of a Medaka Fish Embryo (Oryza latypes) ............................................. 29

3.3 Preparation of a Fly Pupa (Drosophila melanogaster) ............................................... 31

3.4 Preparation of a Plant Root (Arabidopsis thaliana) ................................................... 32

3.5 Imaging Cell Cysts in an Extracellular Matrix Gel ...................................................... 33

3.6 Immunostaining and Preparation of MDCK Cell Cysts .............................................. 34

3.7 Preparation of a Whole Mount of a Mosquito (Anopheles gambiae) ...................... 35

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4 TIPS, TROUBLESHOOTING AND ADDITIONAL INFORMATION ........................ 37

4.1 Tips .............................................................................................................................. 37

4.2 Troubleshooting .......................................................................................................... 37

4.3 Suggested Additional Sources of Information .......................................................... 40

4.4 References and Further Reading ................................................................................ 41

5 INDEX ................................................................................................................. 44

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CHAPTER 2 - SAMPLE PREPARATION Lightsheet Z.1 Introduction Carl Zeiss

1 Introduction

This section describes theoretical and practical aspects of sample preparation for Light Sheet-based Fluorescence Microscopy (LSFM). We present general rules for sample handling and mounting, as well as guidelines with respect to the best preparative technique to use, taking into account sample type, structure and properties. Step by step protocols and recommended materials for Lightsheet Z.1 samples are included. These protocols cover sample preparation ranging from micrometer-sized fluorescent beads to millimeter-sized insects, providing detailed information relating to preparation and observation techniques. Finally, this section identifies the main artifacts and problems that can result from the preparation techniques.

A microscope generally performs best on suitable samples, and when the samples are optimally prepared for the imaging method and microscope type. In Light Sheet Fluorescence Microscopy (LSFM), the sample is commonly mounted in a liquid filled chamber and can be rotated easily. It is scanned through a sheet of light which illuminates the focal plane of a perpendicularly mounted objective lens. The resulting image of an optical section is observed through the objective and is usually detected on a camera-based detector. Since the geometry of the optical beam paths and the optics differ significantly from the conventional inverted and upright compound microscopes, the sample mounting protocols also differ significantly.

If the sample is perfectly transparent, like a block of 1 % agarose with beads inside, the light sheet can penetrate deeply and does not change its properties and shape along the illumination axis. Also, the fluorescent signal can reach the detector unperturbed by scattering effects in the specimen or hydrogel. However, if the sample is slightly opaque and diffracts or scatters the light sheet heavily (lipids, lipid vesicles or dense collagen fiber arrays that scatter light strongly) then the well-defined shape and thickness of the sectioning light sheet degrades along the illumination axis. In a second effect, the detected image from a well illuminated sample might still be degraded by such a poorly transparent sample. These effects can contribute independently to the final image quality of a LSFM and can be minimized or worked around by careful sample positioning in the microscope as well as by an optimized sample preparation protocol.

Ultimately, a fully opaque sample that can completely block the penetration of light and a light sheet (insect cuticular structures, bones…) will limit the imaging capabilities of Light Sheet Fluorescence Microscopes and the Lightsheet Z.1 to its surface.

Furthermore, the image quality in Fluorescence Microscopy in general – and in LSFM in particular - is not only determined by sample transparency that can be optimized by choosing a suitable model (transparent fish like Medaka), suitable growth conditions (no phenol red in the growth media to avoid autofluorescence) or, potentially, a clearing treatment (not suitable for the Lightsheet Z.1). It is also important to have a homogenous signal from a homogeneously labelled sample. Antibodies, for example, are rather large molecules that cannot penetrate deeply into tissue so it is difficult to image a complete juvenile fish after antibody as only the first 50 µm to 100 µm will be labelled, the interior showing reduced signal levels due to the limited diffusion of the antibody.

Samples must be carefully considered when using LSFMs such as the Lightsheet Z.1 as well as the label or dye used must be carefully chosen. In planning an experiment, it should be kept in mind that most labelling and imaging protocols have been developed for thin specimens and therefore many aspects are not adapted to imaging larger samples such as embryos, tissue slices or complete mosquitoes.

Many organisms have been imaged using Light Sheet Fluorescence Microscopy (Table 1) and you may want to read further specific papers to clarify sample preparation issues.

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Table 1

Topic Subtopic Sample/Model

Organism

Physics

Biochemistry

Microbiology

Cell biology

Plant Biology

Developmental Biology

Physiology

Technical set up of MISERB

Structured illumination

Light Sheet Characteristics

Image formation

Image View fusion

Laser Microsurgery

Microtubule dynamic instability

mRNA nuclear export

Heterochromatin dynamics

Imaging of engineered gene expression

Marine microbiology Various bacteria, protozoa etc. LSFM Fuchs et al, 2002

Adaptive optics to improve imaging performance

Intracellular imaging

Nuclear protein localisation

Imaging large living samples

Live imaging of root growth

Consecutive imaging of vertically growing root

Imaging of developing organs

Embryogenesis visualisation

Zebrafish development

Cell identity lineaging and neurodevelopmental imaging

Gene Expression: hour glass model verification

Middle ear structure

3D reconstruction of inner ear

Brain in vivo imaging

3D reconstruction for morphological analysis

Scan of whole brain

Neural network imaging

Sectioning of thick tissues

Imaging neuronal activity

Imaging of immunolabelled receptors

Optical sectioning

Fluorescent beads

Mouse cochlea

Fluorescent beads

Caenorhabditis elegans

live sea urchin embryo, live Danio rerio embryo

In vitro microtubules

Chironomus tentans Salivary Glands

MCDK cells, Drosophila

melanogaster

Drosophila melanogaster

Tumour spheroids

Mammalian cell organelles

Cellular spheroids

MCDK cell cysts

Arabidopsis thaliana

Danio rerio heart valve

Drosophila embryo

Danio rerio

Caenorhabditis elegans

Drosophila melanogaster

Plastic Phantom Structure

Cavia porcellus

Microspheres

Bast's valve

Mouse brain

Mouse cochlea/zebrafish inner ear, brain/ rat brain

Mouse vomeronasal cells

Mouse

Meriones unguiculatus

cochlea, Hippocampus reidi head, Xenopus laevis

Technique/ LSFM

implementation

MISERB

sTSLIM

SPIM

DSLM

LSFM

SPIM

LSFM (FCS)

SPIM

waoSPIM

Bessel beam plane illumination

SPIM

DSLM

SPIM

mSPIM

iSPIM

SPIM

(HR) OPFOS

OPFOS; LSFM

miniSPIM

OPFOS

LSFM

Ultramicroscope

TSLIM

OCPI

SPIM

OPFOS

Reference

Fahrback et al, 2010

Schroter et al, 2011

Ritter et al, 2008

Olarte et al, 2012

Rubio-Guivernau et al, 2012

Engelbrecht et al, 2007

Keller et al, 2008

Siebrasse et al, 2012

Capoulade, 2011

Ejsmont et al, 2009

Jorand et al, 2012

Planchon et al, 2011

Zanacchi et al, 2011

Verveer et al, 2007

Maizel et al, 2011

Sena et al, 2011

Scherz et al, 2008

Huisken et al, 2004

Kaufmann et al, 2012

Wu et al, 2011

Kalinka et al, 2010

Buytaert et al, 2007

Hofman et al, 2009

Engelbrecht et al, 2010

Hofman et al, 2007

Mertz and Kim, 2010

Dodt et al, 2007

Santi et al, 2009

Holekamp et al, 2008

Klohs et al, 2008

Buytaert et al, 2012

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Topic Subtopic Sample/Model

Organism

Large organism general biology

Whole organism 3D reconstruction

Imaging copepod gut contents

Whole organism 3D reconstruction

Ormia ochracea; Emblemasoma auditrix

Drosophila melanogaster

Calanus pacificus

Thermocyclops consimilis

Technique/ LSFM

implementation

LSP

Ultramicroscope

PLIF

LSFM

Reference

Huber et al, 2001

Jahrling et al, 2010

Jaffe et al, 2009

Boistel et al. 2011

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2 Sample Mounting for LSFM

For technical description of the sample chamber, incubation hardware and sample holder of the

Lightsheet Z.1, you should refer to CHAPTER 1 HARDWARE.

The Lightsheet Z.1 system is optimized for gel embedded samples. The sample chamber must be

filled with a watery solution (refractive index of 1.33) at all times, to ensure optimal image quality.

The Lightsheet Z.1 is not designed for the use with clearing reagents. The Lightsheet Z.1 is built for

aqueous media. Also the sample chamber and the system cavity for the sample chamber and sample holder are not compatible with aggressive chemicals.

2.1 The perfect Sample for LSFM

Prior to observing a sample using a microscope, a preparation step is usually necessary. The sample properties and the microscope characteristics provide guidelines and also limitations to sample preparation and imaging. The classic method of mounting an object for microscopy requires the use of a slide and coverslip that in turn limits access to the sample from one side (Fig. 1/A and B). The sample is not touching the objective lens and the number of refractive barriers is at least two (coverslip, mounting medium) and can increase further if immersion oil is needed. The depth of the field of view is dependent on the type of objective lens and the sample properties, and will deteriorate with the thickness of the sample.

Light Sheet-based Fluorescence Microscopy (LSFM) utilizes illumination along an axis that is perpendicular to the detection axis (Fig. 1/C). Moreover, it usually allows sample rotation to record multiple image stacks that are acquired independently along different directions. To allow the high level of sample mobility required for such "Multiview" imaging, the sample is inserted in a sample holder from above. The sample holder is hanging in and linked to an x, y, z, and alpha positioning motor stage, allowing complete three dimensional translations and free rotation around an axis parallel to gravity. This configuration leaves the illumination and the detection paths completely open but requires the preparation of a sample that can be held from above or below in a medium-filled chamber. Such geometry goes hand in hand with the convenient use of water dipping or air objective lenses. As mentioned already in the introduction, another important aspect of sample preparation is the transparency of the specimen, especially when imaging large objects. Ideally, the light sheet penetrates as deeply as possible into the sample. Any obstacle or opaque medium will limit the light sheet penetration depth, generating shadows that will be read out as artifacts on the final image.

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Fig. 1 Relations between the sample and the objective in microscopy.

Samples are traditionally isolated from the objective by a glass coverslip (A and B) limiting access to one side only. (C). In LSFM, the illumination is positioned at 90° compared to the detection axis and can be set up in a sideways geometry ("horizontal microscope").

In LSFM, the sample is usually imaged in a water-based buffer. Generally, it can be kept dry and imaged in air but this has extensive limitations like diffraction due to the significant jump in refractive index from air to the sample material. This has several consequences for sample preparation. First, the refractive index of the mounting medium should be close to that of the sample buffer. The mounting medium should not scatter the illumination or the detection light. Second, the mounting medium should not dissolve in water. Third, its diffusive properties should be close to those of water/medium. Fourth, the medium should be non-toxic for live samples. Fifth, the medium should be flexible to allow the sample to develop. Finally, it should not change its mechanical properties during a period of observation (72 hrs and more).

The following part of this section will deal with sample as a general term but we have devised them in four main classes (Fig. 2) and you can check their size relationship (Fig. 2) and keep that in mind as different samples of different sizes will mean different sample preparation approaches and handling.

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Fig. 2 Different sample types.

(A) very large objects (cm), (B) large objects (mm), (C) medium size samples (100 µm) and small samples (10 µm) (D). Each is represented in relation to the following one to allow size comparison.

2.2 Holding the Sample

In LSFM, the detection axis is at 90° from the illumination axis. There are two main approaches to design such an optical configuration: horizontal or vertical, with respect to the detection axis. In both cases, the sample must be positioned at the intersection between the two axes in order to be observed. The Lightsheet Z.1 is a horizontal LSFM implementation and so the sample is presented from above, hanging along the gravitation axis to be scanned through the light sheet in order to acquire stacks of optical section images. Several possibilities exist to hold the sample in such an optical configuration.

In a vertical configuration, the simplest way is to place the sample on a slide or a cuvette filled with medium underneath the objective (Dodt et al., 2007), alternatively the sample can be embedded in a gel rod that can be rotated. In a horizontal configuration, like the Lightsheet Z.1, the sample can be either embedded in a stiff gel (Fig. 3/A) (Huisken et al., 2004) hooked and positioned in front of the objective (Fig. 3/B) (Engelbrecht et al., 2007), placed in a container (Fig. 3/C) (Engelbrecht et al., 2007, Kaufmann et al., 2012, Pampaloni et al., 2007) or placed on a slide and positioned at a 45° angle (Fig. 3/D). Alternatively, some investigators are using a system presenting the sample from underneath for better stability (Huber et al., 2001).

The Lightsheet Z.1 is not designed to support mounting on a coverslip (Fig. 3/D).

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Fig. 3 Sample positioning in LSFM.

The sample can be held in front of the objective (A) embedded in gel, (B) by a clip, (C) in a container, or on a coverslip (D; Note: the Lightsheet Z.1 is not designed to support this way of mounting the sample). (E to H) show an eye bird view of the mounting (A to D).

Every mounting technique has some advantages and disadvantages. Here, we would like to mention one important parameter: the position of the sample relative to the objective lens. Gel embedding (Fig. 3/A and E) is usually safe but the capillary that holds the gel can potentially touch the detection objective. Such collisions are even more likely for hook (Fig. 3/B and F) and coverslip (Fig. 3/D and H) mountings. It is important to remember that in the Lightsheet Z.1 the sample can interact with the detection lens as well as the walls of the sample chamber and this could affect the imaging process.

One of the important advantages of the LSFM optics geometry is that it allows so-called Multiview imaging. In this case the sample must be mounted to support the required positioning. One approach used in Lightsheet Z.1 to support this experimental paradigm is to place the object in a gel rod that can be rotated (Fig. 3/A and E) in front of the objective. The hydrogel cylinder must be sufficiently stable to avoid movement during rotation. Typical preparation protocols use 0.8 % to 1.0 % agarose (see below in this section) to take this into account. The following sections will address the four main types of sample preparation that can be used: embedding, hanging, enclosing or flattening.

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2.2.1 Embedded Samples

Embedding objects in plastic materials is a routine procedure widely used in the preparation of samples for electron microscopy. In the case of sample preparation for LSFM however, the immobilization of hydrated biological materials must not impair biological activity. It is necessary to keep the object we wish to observe in a perfect condition.

Fig. 4 Embedded samples.

Large samples, such as an adult Drosophila melanogaster, can be embedded in a large gel tube or a cut 1 ml syringe (A), intermediate size samples, such a Medaka or Zebrafish embryo can be prepared by using either a cut 1 ml plastic syringe (see also Fig. 5) or a glass capillary (C and D) and small samples such as Drosophila melanogaster embryos or early stage cell clusters can be prepared using a smaller capillary (D).

In the case of LSFM, it is also necessary to contain the sample in such a way that it can be positioned and rotated in front of the objective. Furthermore, transparency of the mounting medium is essential to allow imaging. A basic technique of mounting objects for the LSFM is to shape them into a cylinder of gel (for example agarose, see also section 2.3.4 Gels and Polymers) that can then be mounted on a dedicated holder. The Lightsheet Z.1 package provides four capillaries sizes adapted to the sample holder to embed objects of various sizes. The special sample holder of the Lightsheet Z.1 adapts to hold these capillaries for precise positioning (translation and rotation) of the cylinder-shaped object for observation through the detection optics. The used gel such as agarose behaves like mechanically stabilized water, supporting the object. It can be easily molded and the gel chosen should have an optical (refraction) index close to that of water. The object can be any size, as the gel can be molded accordingly (Fig. 4/A to D). The various gelling agents and polymers that can be used are discussed in greater detail in section 2.3 Materials and Equipment (see below).

The preparation of embedded samples requires a container suitable for molding the gel. The simplest approach is to use any cylinder with a tight-fitting plunger to pump the molten gel into it and let it polymerize inside before pushing it out. The cylinder can be a syringe, a capillary or even a pipette.

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CHAPTER 2 - SAMPLE PREPARATION Lightsheet Z.1 Sample Mounting for LSFM Carl Zeiss

In the case of Lightsheet Z.1, the Sample Starter Kit provides four types of color coded capillaries and a specific sample mounting device with color coded sleeves to fit each type of capillary perfectly to the sample holder. Moreover, a syringe sample holder is provided.

Below the preparation of a syringe for large sample and a basic protocol for capillary mounting are discussed.

An example of how to tailor a syringe for Lightsheet Z.1 sample preparation is illustrated in Fig. 5. The tip of the syringe is cut off to create an even cylinder, and the gel solution is pumped in using the plunger. The sample is then positioned precisely within the gel.

After the gel has polymerized, the plunger is used to push out the specimen prior to imaging.

Imaging is not done through the syringe or capillary, since this would impair the image quality due to the optical properties of the material.

Fig. 5 Preparing a sample embedding container with a syringe.

Many tubular objects can be used to make embedded samples. A simple technique is illustrated here. The tip of a syringe is cut away (A, B), agarose can then be easily pumped in using the plunger (C), and the sample can be positioned within the agarose tube (D). After polymerization the sample can be pushed out of the syringe for imaging (E).

For smaller samples, a capillary can be used as a sample embedding container. There are several commercial companies that provide glass capillaries with specific Teflon plunger. The Lightsheet Z.1 sample preparation kit comes with four sizes of capillaries and their specific plunger for this purpose.

Make sure you use the right capillary for your sample. The sample size should not be more than 2/3

of the final agarose diameter and no less than 1/3. You should also ensure that you use the right plunger for you particular capillary. Finally, the Teflon plunger should be handled carefully and checked regularly for integrity to avoid leaks that will lead to sliding of the gel rod.

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The important points to consider are that the materials used do not interfere with the gel, the object to image or the sample preparation (chemical compatibility, melting point, transparency etc.), it must be easily prepared or easily purchased, it must be compatible to the LSFM sample holder as well as the x, y, z stage, it must fit to the sample chamber and should not cause damage to the objective lens once rotated or moved. It is also reasonable to consider reusable sample holders to limit waste. We have found that home-made sample embedding container using 1 ml syringes (BD Biosciences, Braun, Terumo or your local laboratory plastic ware supplier), 1 ml plastic pipettes (see your local laboratory plastic ware supplier such as Falcon or VWR), and glass capillaries (Brand, Sutter Instruments or check your local glassware specialist, see also section 4.3 Suggested Additional Sources of Information) (Fig. 6) are particularly effective. The plungers usually come with the cylinders or can be made using metal rods, plastic or metal wires of an appropriate diameter.

Fig. 6 Preparing a sample embedding container with a capillary.

A glass capillary can be used. For the Lightsheet Z.1 they come from the manufacturer (Brand) in just the right length (A). Other capillaries can be cut to an appropriate length (B), the agarose with the object can be easily pumped in using the suitable plungers with Teflon tips (C). If no such plunger is available it can be made, for example, from a piece of electrical wire (D). To avoid leakage, such a plunger can be sealed with nail polish (E) once the sample is pushed out. The sample can then be imaged (F).

Once the sample embedding container is prepared, the sample preparation can begin. The first step consists of preparing the supporting agent at a suitable concentration and temperature. The gelling agent is usually a 0,7 to 1 % solution of low melting agarose in water or PBS, depending of the sample to be embedded (fixed, living, sensitivity to osmotic pressure etc.). If the sample needs to be maintained in a drop of solution or contains water or buffer it is advisable to use a higher concentration of agarose to obtain a final concentration of 1 % once the sample is embedded. The use of low melting-point agarose is recommended (Roth, n° 6351.1) as its melting temperature is only approx. 60 °C and it can be maintained liquid at just above 37 °C prior to embedding.

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There are two principal methods of embedding an object. The first is to directly mix the object with the agarose then pump it into the sample embedding cylinder. This is a convenient way of embedding very small objects such as pollen grains (Swoger et al., 2007); yeast (Taxis et al., 2006) or cell clusters (Pampaloni et al., 2007) or even large objects like fish embryos. The action of pumping in the sample with the agarose results in a self-alignment of the specimen within the tube (Fig. 7/A, E and F).

Fig. 7 Basic principles of sample embedding.

A cylinder with a suitable plunger is used as a mounting device (A). The 1 % low melting-point agarose is melted, then brought to 37 °C, then pumped into the cylinder. (B) The object is then introduced to the agarose with a needle or forceps. (C) Once solidified, the embedded sample can be pushed out and imaged (D). Alternatively, the object, devoid of water, or other solution, is added to a solution of 1 % low melting-point agarose at just above gelling temperature (typically 40 °C) and sucked into the cylinder (E) and then allowed to polymerize. The embedded sample can then be pushed out and imaged (F).

The second method is to fill the sample embedding container with the gelling agent, then to place the object within the gel using a needle or forceps (Fig. 7/A, B, C and D). This approach is more suitable for those samples that cannot be easily aligned using the first technique.

In some cases, it may still be challenging to align the specimen in the most suitable way for imaging. The orientation of the sample must then be optimized, so that interesting details are facing the surface of the agarose cylinder with as little material as possible in the optical path. One solution is to fill a syringe with agarose and allow it to cool until it solidifies. The agarose is then pushed out of the syringe (Fig. 8/A and B). A small V-shaped groove can be cut into the gel and the sample then positioned in the V-groove.

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Fig. 8 Aligning an embedded sample.

The sample can be aligned in a particular orientation to allow the details of interest to be close to the outer surface of the agarose. The solidified agarose is pushed out the syringe a few millimeters and a small v-groove is cut into the cylinder to take up the sample (A). The sample is placed into the v-groove (B). The sample on the agarose is pulled back into the syringe and more agarose is added (C). After the cylinder has completely solidified the sample is pushed out of the syringe allowing free sight on to the sample (D). The same approach can be used to carve a central tunnel in the middle of the agarose to align the sample along the agarose tube axis.

The gel can be cut into various shapes depending on the needs (cylinder, hole). Afterwards the gel with the specimen is pulled back into the syringe and is covered with more molten agarose. The agarose is allowed to cool and solidify, this time period can be shortened by cooling the whole sample. For example the housing of the sample can be rinsed with cold water, although care must be taken to ensure that the polymer does not come into contact with the water, otherwise the cooling agarose would become diluted and lose its stability necessary for holding the sample. After polymerization, the sample is ready for imaging.

2.2.2 Hanging Samples

The Lightsheet Z.1 is optimized for gel embedding samples The sample chamber must be filled with

a watery solution (refractive index of 1.33) at all times, to ensure optimal image quality.

This mounting technique can also be used, but will require some initial adaptations to the sample

holder.

An intuitive way of imaging an object is to simply take it as it is and place it in front of an objective. In an LSFM, this can be done by hanging the object in front of the objective where the axis of rotation and gravity are parallel. This can be achieved using a simple hook made of glass, stainless steel or plastic (Fig. 9/A). This mounting technique can be used for large samples such as organs (for example the brain) or complete organisms (insect, fish). One main drawback is the fact that the hook will partially damage the object and may also interfere with the field of view.

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The Lightsheet Z.1 has a maximum Field of View of approx. 2.5 mm (depending on zoom settings).

This and the dimensions of the sample chamber might limit the size of the sample.

Fig. 9 Different ways of hanging a sample.

Samples can be either hooked or deposited on a bent glass capillary (A, D), glued to a rod or capillary (B), or clamped on a syringe tip using the plunger and the syringe body as a holder (C).

Interestingly, such a hook can also be tailored to mount small objects embedded in agarose. The drop of agarose is more stable as it is closely held by the hook. This is very important when imaging at very high magnification (100x). The hanging method has been successfully used for imaging single Saccharomyces cerevisae (Taxis et al., 2006) and holding fish fins during laser nanosurgery (Engelbrecht et al., 2007).

2.2.3 Enclosed Samples

The Lightsheet Z.1 is optimized for gel embedding samples The sample chamber must be filled with

a watery solution (refractive index of 1.33) at all times, to ensure optimal image quality.

This mounting technique can also be used, but will require some initial adaptations to the sample

holder.

The last important technique of holding samples to be mentioned in this section is to create a container that can hold the object in front of the objective lens. This technique is particularly suitable for specimens that should not be embedded (for example due to temperature, physical constraints etc.) or that need to be constantly maintained in a specific buffer (for example in vitro assays, or living cells). The container must be suitable for LSFM imaging. It must be basically transparent and be suitable for the object but also for the imaging chamber and the sample holder. It can be hooked or clipped using specific holders.

There are two main methods of generating such containers, using gelling agent to shape out a container (Fig. 10/A) or using polymers such as PTFE (Polytetrafluorethylen, Teflon) or FPE (Fluorinated Ethylene Propylene) to make it (Fig. 10/B and C).

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Fig. 10 Enclosed chambers for LSFM.

Incubation chambers can be made by molding an agarose beaker that can be mounted on a simple plastic holder and loaded with the sample prior to imaging (A). Another solution is to create a chamber with a specific polymer with a refractive index close to water (for example PFTE or FPE) using heat or glue to seal the chamber to the needed size and volume, and attach it to a suitable holder (syringe, capillary etc…) (B). A PDMS, FPE tube or glass chamber can also be considered (C).

The container can be easily molded using a gelling agent specifically chosen for its stiffness and transparency. The custom-made molding system is made from a syringe where the plunger has been modified to hold a cylinder of smaller diameter. This system allows tailoring of the size of the container wall and is easy to use for molding (Fig. 11/A). The plunger is pulled into the syringe body and filled with molten gelling agent. The plunger is further pulled to create the bottom part of the container (Fig. 11/B and C). Alternatively, the system can be used to generate a hollow tube that can be subsequently sealed (Fig. 11/D). The gelling agent is left to polymerize and then the tube is removed from the modified plunger. The containers can be used directly or kept in a water-based buffer for later use. The gelling agent must be transparent, and although the use of agarose is possible, the concentration will depend on the size of the container walls and the inside chamber. It is recommended to use a higher concentration of gelling agent to ensure the stability of the container. We have used a 1ml syringe as a molding system and a concentration of 1.5 % agarose for the container molding. The stability is good and the degradation of the optical path is minimal. Higher agarose concentrations may generate aberrations.

Fig. 11 Making and mounting an agarose incubation chamber.

A modified syringe plunger is made by inserting a smaller diameter cylinder on to the plunger (A, B). The tube is molded by simply pouring the molten agent into this device (C). Once removed, the open end of the container (D) can be closed with agarose (E).

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Another possibility is to use a polymer to make the chamber. The polymer, similar to the gelling agent, must be transparent or at least have an optical index as close as possible to water or the buffer used during the experiment. The polymer is usually used as a sheet that can be formed as required. The other possibility is to approach commercial manufacturers to make polymer chambers at the specific sizes and lengths required.

Fusing polymer sheets can be done using a welding iron with controlled temperature or a welding device use for melting together plastic bags. As described in Fig. 12, the polymer foil is folded to an appropriate size. This can be made easier by using a guide or template, in this case a micropipette. The polymer is fused together. The tube generated is finally fused together on the other side to make a complete container. The polymer chamber can be easily mounted on the LSFM by using a clip, a slotted metal capillary or glued to a micropipette. However, the last two options have the disadvantage of partially obscuring the field of view.

Fig. 12 Making and mounting an incubation polymer foil chamber.

A piece of polymer foil (A) is folded and either heat- or glue-sealed to generate a tube of a predefined size (B). Excess foil can be removed or used to glue the tube on to a specific holder (capillary, thread, metal rod…) (D). One side of the tube can be then glued or heat-sealed to close the chamber (C). The polymer used must be suitable for microscopy and easy to seal. The chamber can be glued to a support, held by forceps, or inserted into a slit rod.

This technique has been successfully used to image living cells (Engelbrecht et al., 2007) and cell clusters (Pampaloni et al., 2007).

2.2.4 FEP Tubing

More recently, the availability of Fluorinated Ethylene Propylene (FEP) tube of different diameters has been successfully used for long term imaging of Zebrafish embryos (Kaufman et al., 2012). Here, we refer only to the paper by Anna Kaufmann, Michaela Mickoleit, Michael Weber and Jan Huisken in Development 139, 3242-3247 (2012) ("Multilayer mounting enables long-term imaging of zebrafish development in a light sheet microscope") and emphasize the fact that the mounting method described in this article is fully compatible with the Lightsheet Z.1.

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2.3 Materials and Equipment

This section gives an overview on the materials and equipment for sample mounting and the sample chambers of the Lightsheet Z.1. The generalization of the concept is also mentioned.

2.3.1 Sample Chambers

In the Lightsheet Z.1, the sample is positioned within a chamber containing an aqueous solution. This chamber is tailored with O-rings to tightly fit the detection optics and avoid leakage. The upper part of the chamber is open to allow introduction of the sample. The bottom can be equipped with a Peltier Block or a Heatingblock (optional incubation). The remaining three sides are made in such a way that glass coverslips can be fixed allowing entrance of the light sheets from two sides and observation of the object by the user during the different steps of imaging using the appropriate software feature. The original chamber is made of medical steel, however, depending on the buffer used (salt, pH etc.), the experiment being performed (time lapse, live cell imaging etc.), there might be a need for more specific chambers. Carl Zeiss provides the technical drawing of the sample chamber for Lightsheet Z.1 so that users can develop their specific sample chamber manual for further information.

. You can refer to the sample chamber section of this

When designing a chamber for your particular application you must take into account the following points:

Transparency: user visual access, light sheet entry and exit routes, the distance of the coverslips for

the light sheet – and the water filled space in between - are a crucial measure in the optics calculation of the Lightsheet Z.1 system. To ensure the functionality of the system these have to be maintained when a custom made chamber is designed.

− Temperature control: heating devices, cooling devices

− Volume: size of the sample, buffer used (cost), drug treatment (cost)…

− Fitting: objective, illumination position, stage, heaters…

− Material: buffer, heater, sterilization, UV protection…

− Flow: flow entry and exit

− Size

− Cost

Carl Zeiss Microscopy GmbH (hereinafter "we") hereby informs you that we will warrant the specified and agreed performance of the

Lightsheet Z.1 system only if sample chambers are applied and used that either are delivered or explicitly approved by us.

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1 Front system cavity door 6 Sample chamber 2 Upper sample opening 7 Guide rails 3 Upper system cavity door 8 Connections for incubation 4 Securing screw 9 Hose and syringe 5 Sample chamber grip

/ sample chamber mount

Fig. 13 Removing and inserting the sample chamber.

The chamber has five entry points allowing the positioning of the objective, the sample holder, the light sheet and the observation by the user. Heated chamber. The chamber can be equipped (optional) with a Peltier Block that can be tuned according to needs or a Heatingblock. For further details on the sample chamber handling, accessories, its cleaning and assembly please read the corresponding CHAPTERS of this manual.

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2.3.2 Molding and Mounting Supports

As described previously, there are several options to prepare a sample and therefore several options to manipulate and mount it. Initially, readily available products found in cell biology laboratories: syringes, capillaries or pipettes were used to mount samples. These components are all commercially available, cheap and convenient for LSFM sample preparation. However, they still need to be prepared for the specific needs. Plastic syringes exist in various sizes (0.2 ml, 0.3 ml, 0.5 ml, or 1 ml) and have tight plungers that easily allow pumping and movement of the agarose rod used to embed the sample.

They can also be used to hang the sample by effectively using the plunger and syringe body as forceps. The sample holder disc for syringes (Fig. 14/I and K) provided should be used in this case. Moreover, they can be purchased sterile for single use applications.

In the case of Lightsheet Z.1, the sample kit is provided with four types of color coded capillaries with matching plungers (Fig. 14/A and B) and color coded sleeves to fit perfectly each type of capillary to the sample holder (Fig. 14/C). The typical protocol of sample mounting is a two-step process of choosing carefully your sample mounting system based on your sample (size, agarose/sample ratio…) then to assemble it (e.g. plunger+tip+capillary) beforehand. Prepare it (e.g. sample + agarose) and insert it in the Lightsheet Z.1 upper sample opening (Fig. 14/G and H).

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Fig. 14 Sample mounting accessories as part of the sample chamber and holder starter kit.

A. Capillaries (inner diameter of capillary size 1 / ~0.68 mm, size 2 / ~1 mm, size 3 / ~1.5 mm, size 4 /

~2.15 mm) B. Specific plungers and Teflon tips for each capillary. C. Specific color coded sleeves to adapt each capillary to the sample holder (F). D. Sample holder stem for capillaries, clamp screw, ejection tool. E. Sample holder disc for capillaries F. Sample Holder diagram showing the capillary, the stem and disc of the sample holder. G and H. Sample holder handling and insertion in the Lightsheet Z.1. I. Sample holder disc for syringes, adapter ring. K. syringe (1 ml).

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Capillaries are made of glass. They can break. They are also sliding when wet. Please handle

Carl Zeiss Sample Mounting for LSFM Lightsheet Z.1

A few points must be taken into account when choosing a particular mount:

− Compatibility. This is a crucial issue. The mount must be compatible with the object you want to image (chemistry, temperature etc.), but it must also be compatible with the stage holder.

− Stability (mechanical, optical, chemical).

− Tightness. In the case of embedded samples, once the gel has solidified, the cylinder of gelling

agent is pushed through the capillary out of the distal end by a plunger fitting into the capillary. The system must be air tight to avoid air entry leading to a displacement of the gel rod. The plunger can be sealed with a drop of wax, acrylamide or nail polish, i.e. anything that prevents the plunger and hence the agarose containing the sample from moving.

− Cost.

2.3.3 Sample Holder

For technical description of the sample chamber, hearting devices and sample holder, you can refer to CHAPTER 1 HARDWARE.

Once the specimen is prepared and properly labeled, it is ready to be imaged. While in conventional imaging there is a suitable platform on which to place the glass slide or the chamber, in LSFM the object must be held from above via the sample holder. Depending on the size of the sample there are two different types of sample holders available: sample holder for capillaries and syringes (Fig. 14/C-F and I). Always use the minimal cylinder diameter necessary for your specimen size to avoid excessive amounts of agarose.

The largest sample holder has been designed to accommodate a 1ml syringe that can be inserted from the top with a plunger that can be operated once the sample holder is mounted on the stage. Once inserted, the syringe is perfectly fitted to the sample holder as the two flaps used for injection fit the upper part of the holder. In this way the object support is well maintained, an essential issue for imaging and multiview imaging as the object is moved through the light sheet by the stage.

Capillaries have been extensively used to image small embedded objects, as hooks for very large objects, and as support for enclosed objects, so the capillary has become commonly used for LSFM sample embedding.

them with care and dispose them properly.

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2.3.4 Gels and Polymers

Gelling agents are commonly used for preparing semi-solid or solid tissue culture media. Gels provide support to tissues growing in static conditions. The gelling agent usually has several properties. In particular, it does not react with media constituents, is not digested by enzymes, and remains stable at all incubation temperatures. Gelling agents are very versatile and useful tools in LSFM as they allow easier sample preparation. This section will present in more detail the properties, advantages and disadvantages of two well-described gelling agents and provides an additional list of gelling agents.

Agarose

Agarose is a complex carbohydrate polymer material, generally extracted from seaweed. It is used in chromatography and electrophoresis as a medium through which a substance can be analyzed by separating it into its components. The molecules are extremely water-soluble due to their large number of hydroxy groups, and solutions tend to be low-melting point aqueous gels. A wide range of different agaroses, of varying molecular weights and properties are commercially available. These include low melting types, (for example, Agarose Type VII, low melting temperature: gelling temperature below 30 °C, melting temperature above 65 °C) which can be used if the sample is sensitive to high temperatures. Interestingly, the refractive index of the low melting type is lower than that of normal agarose. However, to obtain the same strength, a higher concentration needs to be used. With a concentration of 1 % (w/w) the low melting point agarose has the same stability as a 0.5 % agarose (normal). The refractive index at this concentration is still lower than that of normal agarose, minimizing distortions when imaging. In our laboratory, we preferentially work with agarose as it is easy to handle, has good optical properties and is not expensive.

Gelrite

Gelrite gellan gum is a self-gelling hydrocolloid that forms rigid, brittle, transparent gels in the presence of soluble salts. Chemically, it is a polysaccharide comprised of uronic acid, rhamnose, and glucose. It is produced by the bacterial strain S-60 of Pseudomonas elodea. Gelrite is a trademark of Merck and Co, Inc (Rahway, NJ), Kelco Division, USA. One advantage of Gelrite is the lower scattering of light compared to an agarose gel with the same stability. It has a higher index of refraction but less scattering compared to agarose. Gelrite has a consistent batch-to-batch quality due to a stringent control of the fermentation process. Only half the amount of Gelrite is required for the same purpose. It hydrates rapidly and gel setting can be easily controlled. The stability of the gel depends on the concentration of divalent cations

(Mg

, Ca2+) therefore a gel made with Gelrite and pure water is unstable compared to a PBS- (buffer) based gel. Polymerisation is faster compared to agarose, which might be advantageous for some applications. The temperatures for gelling and remelting are similar to that of agarose.

Additional list of gelling agents

− Galactan

− Agar

− Gelatin

− Carrageenan

− Alginate

− Phytagel™

− Agargel™

− Transfergel™

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2.3.5 Hydrogel Preparation

Every gelling agent is prepared following specific protocols that vary widely from supplier to supplier, and from laboratory to laboratory depending on the final application. We will not try to cover every single one of these but rather give a simple protocol that we have been using in our laboratory to prepare embedded samples in agarose as a gelling agent. The preparation is done as follows:

1. Preparing a 1 % low melting agarose gel

Weigh 1 gram of low melting point ("low gelling") agarose ("Agarose Low Melt" (no 6351.1 from "http://www.carlroth.com") and place it in a flask. Add 100 ml of solvent (water, PBS) to the flask. Swirl to mix the solution.

Place the flask in the microwave. Heat above 95 °C until the solution is completely clear and no small floating particles are visible. Do not allow the agarose to boil over as this will affect the final agarose concentration. Swirl the flask frequently to mix the solution, prevent the agarose from burning, and prevent boiling retardation.

Wear heat-protective gloves when handling the flask.

The agarose can be also sterilized for sterile use (cell culture). It is also possible to remove dissolved air bubbles using a vacuum pump.

2. Cooling the gel

Once molten the gel is left to cool to 37 °C (or just above gelling temp - read the material properties sheet) in a water bath or on a heating plate. It is very important, especially for sensitive samples, to ensure that the agarose is at 37 °C before use.

Note: Alternatively, you can aliquot your agarose solution into 1 ml or 2 ml Eppendorf tubes for later use. Label them and store them in a cool and dry place. In this case, each aliquot can be liquefied using a heating block (80 °C – 90 °C) then transferred to a heating block at 37 °C.

3. Using the gel

At this stage follow the examples described in the section dealing with embedded sample preparation. Avoid bubble formation during handling and pipetting as they will impair the embedding process. Work quickly as the low melting point agarose will polymerize rapidly as it was kept at 37 °C close to the gelling temperature.

4. Polymerization of the gel

Let the gel polymerize. Avoid contact with any water-based solution as it will dilute the gelling agent solution. The process of polymerization can be accelerated by cooling down the embedded sample (cold water, fridge…) – but keep in mind that this might affect viability of a living sample.

5. Using the prepared sample

Once fully polymerized, the embedded specimen can be manipulated, but keep in mind that it is a gel and therefore fragile. Avoid any kind of friction, or shock etc. As it is a water-based object it must be kept wet at all times to avoid drying out and damaging the sample. Moreover, many types of gel may change their properties over time (e.g. swelling), and this can result in loosening of the gel in the support. It is therefore important to use your sample as soon as possible and monitor its quality over time if you plan to reuse it.

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2.4 Fixation and Fixatives

Many experimental samples will require fixation prior to imaging. The goal of fixation is to maintain cellular structure as close as possible to the native state. Proper fixation typically facilitates immunohistochemical analyses if desired, and is an important step prior to further processing. Specialized fixation procedures and processing may be required for certain tissues (e.g. bone de-calcification) or preserving specific target antigens.

The processing of most samples begins with fixation to preserve morphology. A fixation method must take into account two things: the preservation of cellular 3-D structure and maintenance of good access to antigenic sites. The goal is to preserve sufficient cellular organization to allow identification of the features of interest, but not to destroy the antigenicity of the target. Fixation is also frequently combined with permeabilization to allow the staining solutions used in later steps access to the cellular interior.

Commonly used histological methods of fixation and permeabilization often consist of treating the cells with solvents, such as methanol. While these methods are rapid-acting precipitating fixatives, they are also good permeabilizing agents, but have one significant negative consequence: cellular shrinkage. The degree of shrinkage may be almost insignificant for monolayers of cells, but will distort tissue samples dramatically. To take full advantage of the three-dimensional reconstruction capability of the LSFM microscope, the use of a fixative that does not destroy in vivo structure and organization is imperative.

It is important to remember that different specimens may require different fixation methods. Testing and optimizing for each new sample type will ensure that the best balance between preservation and labeling is obtained. Fixing and permeabilizing your cells affects the cell morphology and the availability of the antigen you are trying to detect. You may get different results with different reagents, times and concentrations, hence the need for protocol optimization. The distortion of cell morphology is something to bear in mind when interpreting the images.

2.5 Stains and Staining

In LSFM, like in any microscopy technique using fluorescence, the sample can be labeled using specific fluorescent dyes, fluorescent proteins or fluorescently coupled antibodies. Two basic techniques are generally used: direct labeling and indirect labeling. Both labeling methods are suitable for LSF microscopy. Direct labeling consists of using fluorescent proteins, fluorescently labeled primary antibody or a dye that cause the structure of interest to become fluorescent. Advantages of this method include speed and ease of application. A potential disadvantage is lack of sensitivity (low signal intensity). The indirect method involves binding a primary antibody to the epitope of interest, followed by a fluorescently labeled secondary antibody. The main advantage of using this technique is the great amplification of signal possible through an antibody cascade. The disadvantages include increased complexity, the method is more time consuming, and there are often problems with non-specific antibody reactions.

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2.5.1 Choosing a Fluorescent Label

The choice of label depends upon the available equipment on your LSFM set-up (lasers, filters) and the availability of certain fluorescent protein variants, fluorochromes conjugated to required antibodies for use in multiple labeling schemes. In general, the laser lines available dictate which fluorophores or fluorescent proteins can be used. Recent advances in biochemistry have created new families of fluorophores with very favorable signal-to-noise and quantum efficiency (QE) properties. Similarly, many laboratories have developed a wide variety of fluorescent proteins that span the spectra from GFP

Plum.

2.6 Antifading Agents

Fluorescently labeled cells and tissues exhibit a characteristic photobleaching curve in response to excitation by the light. Much of the photobleaching can be attributed to the generation of free radicals. The use of free radical scavengers has been shown to decrease the rate of photobleaching. Common scavengers include n-propyl gallate, p-phenylenediamine and DABCO (1,4-diazobicyclo-(2,2,2)-octane). Live systems have been reported to reduce photobleaching in the presence of vitamin C or Trolox. As the LSFM technology reduces greatly the phototoxicity and photobleaching effects during imaging, we never encounter samples that require the use of antifading agents so far. However, some applications may require the use of radical scavengers during long time imaging of GFP expressing samples as repeated exposure may lead to a regular increase of the free radical contents, which might affect its behavior over time.

2.7 Cleaning, Labelling and Storing Samples

One important point about samples is their handling. In the case of LSFM, all the samples are three dimensional objects that are mounted to be imaged in a chamber containing water based medium. They must then be maintained in a moist environment. Once prepared and prior to imaging, the samples can be held in a filled beaker or Falcon tube filled with the appropriate medium, e.g. water, PBS (Fig. 15). One simple solution we have developed in the laboratory is to use a beaker filled with the right buffer. The samples are maintained by using plasticine on the beaker border. An alternative is to cover the top of the beaker with an aluminum foil and accommodate the sample holders such as the 1 ml syringe by drilling a hole in the foil. This handling technique limits evaporation. More advanced holders can be designed and manufactured according to need. You will find a couple of examples that we have made in our laboratory keeps the samples moisture at all time. They are stable and can be easily move from the laboratory to the microscope as well as stored in the fridge.

to handle various size of sample embedding containers. They include a water tank that

Flood P.M., Kelly R., Gutiérrez-Heredia L. and E.G. Reynaud

School of Biology and Environmental Science, University College Dublin, Belfield, Dublin 4, Dublin, Ireland

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Fig. 15 Supports for sample embedding containers.

Support are used to hold three dimensional objects that cannot be held flat easily. Moreover, embedded samples need to be kept in buffer to avoid gel shrinkage or sample damage. A simple system is to use a beaker filled with PBS and place plasticine on the upper border to support the sample embedding container (A). More elaborate supports can be made using clips of different sizes for holding syringes (B) or even capillaries (C).

As in LSFM there is no need for oil or any specific chemical for imaging, the cleaning of samples is not necessary. However, you can rinse the sample within the capillary or syringe with water or your specific buffer after imaging if the chamber was containing particles, bacteria or other chemicals (dyes, drugs etcetera).

Labeling the samples can be an issue as it can be tricky to mark the name of every sample on the embedding container (capillary, syringe…). A simple marking technique is to use tape roll around the syringe plunger or the capillary. This must not affect the handling of the sample on the microscope. Another approach is to number the sample and to register the detail on a lab book. However, this can be a problem if you store many sets of samples in the same fridge day after day.

Sample preparation techniques usually allow long time storage (paraffin embedding, slides…). As long as a few basic rules are followed (keeping away from light, temperature…) they can be kept up to years. In the case of LSFM, the samples are imaged in a water environment and must be always kept wet, even for long time storage. This can be a challenge. Usually, we keep fixed samples in the fridge using a sample embedding container support and we refill the buffer tank from time to time. However, we never kept samples for more than a month under such conditions. A longer storage possibility is to use a water tight container where the samples are kept with enough water not to dry out. One point to consider is the way the sample was prepared. Embedded samples may weaken with time as some gels may not maintain their strength over time at 4 °C. Hooked samples may as well be loosening and fall from their support. It may be better to unhook them and store them in a different type of container.

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3 Specific Examples of Sample Preparation

In order to make this sample preparation section as useful as possible the following pages describe mounting techniques for specific samples, in particular describing the equipment needed, step by step protocols and illustrations based on our own laboratory

Fluorescent beads are used for later Landmark Registration processing of acquired multiview data,

and should be included during embedding of samples of interest. Prepare agarose as described in section 3.1 Preparation of Fluorescent Beads accordingly.

3.1 Preparation of Fluorescent Beads

Samples with fluorescent beads are often used to characterize the imaging properties of a microscope such as the LSFM. Using a reproducible sample is an important tool to calibrate the instrument. This protocol describes how to handle fluorescent beads and to prepare optimal concentrations to image with an LSFM.

experiences.

Equipment and reagents

− Fluorescent beads

− 1 % Low Melting Point (LMP) Agarose in deionised water

− Capillary (Size 4, Blue, #701910, BRAND GmbH)

− Sonicator

− Heating block- Vortex

Method

1. Vortex the bead solution to make a homogeneous dispersion.

2. Dilute a small volume of the bead dispersion in deionized or distilled water to a concentration 100x higher than the one desired for the specimen. Depending on the size of the beads and the magnification required it is first necessary to calculate the bead-agarose ratio (see below).

3. Sonicate the dilution for 5 minutes at maximum power.

4. Prepare a liquid agarose solution of a chosen concentration (0.5 % - 1 %) and cool it down to just above the gelling point (usually 38-40 °C).

5. Mix diluted fluorescent beads with the agarose in ratio 1:100 and vortex the mixture. Use a pipette or a capillary (by sucking in and out the liquid agarose several times) to mix the bead solution and the agarose thoroughly.

6. Insert an appropriate plunger and Teflon tip.

7. Push the plunger through the capillary, so the front end of the plunger is sticking out of the capillary by a bit before entering the liquid agarose and sucking the agarose in. This will avoid air bubble formation at the plunger.

Flood P.M., Kelly R., Gutiérrez-Heredia L. and E.G. Reynaud

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8. Suck in the agarose/beads by pulling the opposite end of the wire/plunger.

9. Let the gel polymerize (approx. 5 minutes) before imaging.

10. Make sure that only a very short part of the agarose block is pulled out of the glass capillary during image acquisition.

11. When multiple views are recorded, it is best to image from the centre of the agarose block.

Beads should be fluorescent in the part of the spectrum you would like to analyze for 1 channel

systems. With the Lightsheet Z.1, a two channel system, one can use one channel for the beads (e.g., red) and one channel for the specimen label (e.g, GFP.) Fluorescent beads covering whole visible spectrum are nowadays easily available from various different suppliers (e.g. Polysciences, Invitrogen, Estapor / Merck etc.). In our case, the density of the fluorescent beads is chosen to end up with several hundred beads in the imaged volume. For example, for a 40x magnification lens the

volume of interest is around (200 µm) beads are shipped as a solution of 5*10 them 1:10

in agarose to have approximately 400 particles in the volume of interest. Having too few of them (less than 100) in the three-dimensional image will give you no or poor processing results, while too many of them (more than 1000) might considerably increase processing time without a significantly improving the final results.

= 8*10-6 ml. If the fluorescent

particles/ml , you have to dilute

Moreover, a gel with sufficient stiffness but minimal impact on the image has to be used to immobilize the beads. 1 % low-melting agarose (Sigma, Type VII) is being routinely applied for lenses with numerical apertures up to 0.8 NA. For 1.0 NA objective lenses and above, a more diluted (e.g. 0.5 %) gel must be used to minimize gel-caused image aberration.

3.2 Preparation of a Medaka Fish Embryo (Oryza latypes)

Embryos have been extensively used for many decades to study developmental mechanisms as well as diseases. They can range from micrometers to centimeters depending on the species used (frog, fish, fly, worm, etc.). This protocol applies to most embryos. The important point is the temperature. The embryo must not be damaged by temperature shock during embedding. Moreover, the embryo should not be constraint by the stiffness of the gel. This may impair its normal development.

Equipment and reagents

− 1.5 % Low Melting Point (LMP) Agarose in E3 (Fish buffer)

− Mesab/Tricaine 0.4 % stock (3-Aminobenzoic Acid Ethyl Ester)

− Capillary (Size 4, Blue, #701910, BRAND GmbH)

− Electrical thread (1,6 mm) or plunger

− Heating block

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Fig. 16 Mounting an Oryza latypes embryo. (A) The embryo is prepared (labelling, drug treatment,

dissected...) (B) The embryo is deposited on the side of the Eppendorf tube and the excess of water is removed with a pipette. (C) The embryo is dropped into the agarose and pumped into the capillary. (D) The embryo can be imaged.

1. Select embryos for imaging, dechorionate. Melt 1.5 % LMP agarose, aliquot 0.5 ml into a 1.5 ml Eppendorf tube. Add 150 ml of Mesab to ensure that the embryos do not move during imaging. Invert the tube to mix and allow agarose to cool to 40º C.

2. Add the embryo to the tube containing agarose using a Pasteur pipette, transferring as little buffer as possible.

Add the embryo to the Eppendorf tube as a drop on the tube wall. If necessary remove the extra

buffer with a yellow tip before dropping the embryo into the agarose.

− Or transfer the embryo to an empty Eppendorf tube, remove all medium and add the liquid agarose.

3. Let the embryo fall to bottom of the Eppendorf tube. Insert a capillary into the tube and suck the embryo into it by pulling out the thread or plunger like a syringe piston.

− When sucking up the agarose, make sure that initially the plunger is sticking out of the capillary within the liquid agarose, to avoid air bubble formation. Furthermore leave some space between the plunger and the sample (see Fig. 16/D)

4. Allow the agarose to harden and place the capillary in a stand in water or PBS.

5. Mount on the capillary sample holder prior to imaging.

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3.3 Preparation of a Fly Pupa (Drosophila melanogaster)

Some type of embryos are hydrophobic once dissected and cannot be mounted using the technique described above as they will float on the agarose and will be impossible to embed. The following protocol is suitable for this type of embryo such as fly embryos or pupas.

Equipment and reagents

− Drosophila melanogaster pupa or embryo

− 1 % Low Melting Point (LMP) Agarose in water or PBS

− Capillary (Size 4, Blue, #701910, BRAND GmbH)

− Heating block (90 °C and 40 °C)

Method

6. Choose a pupa Drosophila melanogaster. Melt 1 % LMP agarose, aliquot 0.5 ml into a 1.5 ml Eppendorf tube. Invert the tube to mix and allow agarose to cool to 40º C.

7. To allow sample preparation the pupa must be submerged in agarose by pouring it directly on top of it in a large drop of molten low melting point agarose.

8. The pupa can then be pumped into a capillary as previously described.

9. The insect can then be imaged.

Fig. 17 Mounting a Drosophila pupa.

(A) The pupa is prepared (labeling, drug treatment, dissected...) (B) The pupa is deposited in a watch glass and covered by melted agarose to embed the hydrophobic pupa. (C) The pupa is then pumped into the capillary. (D) The pupa can be imaged.

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3.4 Preparation of a Plant Root (Arabidopsis thaliana)

Plant research is an important field of investigation using plants as model systems. They are threedimensional objects that are difficult to image fully and are usually dissected and sliced before being imaged and analyzed. This protocol has been used to mount complete young Arabidopsis thaliana plants for imaging root development directly on the microscope.

Equipment and reagents

− 1 % Low Melting Point (LMP) agarose in plant buffer

− 1 ml syringes

− Arabidopsis thaliana seeds

− Heating block

Fig. 18 Mounting an Arabidopsis thaliana root

(A) An Arabidopsis thaliana seed is positioned at the bottom of an agarose cylinder (B) After a few days of development, the root can be seen in the agarose cylinder bottom. (C) The agarose cylinder is pushed out of the syringe for imaging.

Method:

1. Several agarose beakers are prepared as described in the enclosed sample section.

2. Instead of pushing out the plunger to extract the agarose beaker, the plunger is pulled in to the end of the syringe where it can be released leaving the beaker inside the syringe.

You need to make a long walled beaker to avoid inconvenient breakages and leakages that may be

caused by the following steps.

3. A seed of Arabidopsis thaliana is put at the bottom of the beaker.

4. The beakers are kept in the syringe in a humidified and well lit chamber to allow seed germination.

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5. Once the root is visible within the bottom part of the beaker, a normal plunger is inserted in the top part of the syringe, where the open part of the beaker is present.

6. Push down the beaker into the syringe until the root can be seen outside of the syringe cylinder

7. The beaker is mounted in the sample chamber filled with distilled water or plant growth media at room temperature.

As the plant depends on light to grow long-term imaging must take into account the illumination

of the leaves between imaging sessions.

3.5 Imaging Cell Cysts in an Extracellular Matrix Gel

Live imaging of cells has been a major tool in cell biology. For this, cells must be maintained in optimal conditions during the complete time of the experiment. The incubation options for the Lighsheet Z.1 are described in another section of this manual (CHAPTER 1 HARDWARE). However, cells must be mounted in a way that allows them to hang in front of the objective from above. This protocol describes one way of imaging MDCK cells that naturally form cysts when grown in an extracellular matrix.

Equipment and reagents

− MDCK cells grown in an extracellular matrix (Matrigel, ExtraCell etc.)

− 1.5 % Low Melting Point (LMP) Agarose in PBS

− Modified plunger

− Sealing device

− Slitted capillary

− 1 ml syringe

− Capillary holder

− Heating block

− Polytetrafluorethylene foil or FPE tube

Method

1. An agarose beaker or a polymer foil chamber is prepared as previously described in the section

2.2.3 Enclosed Samples.

2. MDCK cysts are grown in an extracellular matrix gel.

3. Cells can be stained at this stage with live markers (nuclear, mitochondrial, lysosomal etc.) before mounting.

4. Cells within the gel are transferred into the chosen chamber (agarose, polymer) and mounted in a 37 °C and CO

chamber on the LSFM using a cut tip to limit shearing damage.

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If cells are grown in a different manner it is possible to mix them with a supporting gel prior to

loading into a chamber. They can also be grown within the gel already present in an incubation chamber. This limits damage, shear and temperature changes during sample preparation and handling.

5. The agarose incubation chamber is mounted on a specific holder. The polymer chamber can be either clipped or glued to a supportive holder.

Eukaryotic cells are highly sensitive to environmental change (temperature, pH, osmotic pressure

etc.). The transfer steps must be rapid and carried out in a sterile manner (wherever possible) especially for long term time lapse experiments. It is important to be gentle and use cut tips and pre-warmed materials at all times, including the sample chamber.

6. Monitor the cell status during imaging to check viability and changes.

3.6 Immunostaining and Preparation of MDCK Cell Cysts

Immunofluorescence allows highlighting of specific proteins or structures using specific antibodies. This protocol is used to perform immunofluorescence on cysts which are three-dimensional cell structures that can be grown in extracellular matrix gel such as collagen.

Equipment and reagents

− 1.5 % Low Melting Point (LMP) agarose in water or PBS

− Capillary (Size 4, Blue, #701999, BRAND GmbH)

− Electrical thread (1.6 mm) or plunger

− 4 % paraformaldehyde solution

− Antibodies (primary and secondary)

− PBS

− Triton X-100

− Bovine Serum Albumin (BSA) or Foetal Calf Serum (FCS)

− Heating block

Method

1. MDCK cell cysts grown in extracellular matrix are collected and centrifuged at 500-1000g to pellet the cysts with the gel.

2. The supernatant is removed and replaced with 4 % paraformaldehyde and incubated for 15 minutes on a wheel or rocker to efficiently mix the gel pellet within the fixative.

3. The gel is pelleted and the supernatant is replaced by 0.1M glycine to quench the paraformaldehyde, and then incubated for 10 minutes.

4. The gel pellet is washed twice with PBS (500-1000 g, 5 minutes).

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5. The pelleted cysts are permeabilized with PBS/1 % Triton X-100 for 10 minutes on a wheel or rocker to efficiently mix the gel pellet.

6. The gel pellet is washed twice with PBS (500-1000 g, 5 minutes).

7. The gel is incubated for 10 minutes in PBS/1 % FCS on a wheel or a rocker to block the extra epitopes and efficiently mix the gel pellet.

8. The gel pellet is incubated with the primary antibodies at the concentration indicated by the supplier, using a wheel or rocker to efficiently mix the gel pellet.

9. The gel pellet is washed twice with PBS (500-1000g, 5 minutes).

10. The gel pellet is incubated with the secondary antibodies at the concentration indicated by the supplier, using a wheel or a rocker to efficiently mix the gel pellet.

11. The gel pellet is washed twice with PBS (500-1000 rpm, 5 minutes).

12. The cysts can be stained at this stage with Hoechst to label the nuclei.

13. The gel is pelleted and as much of the supernatant as possible is removed.

14. The gel pellet is mixed with low melting point agarose, mixed and pumped into a capillary.

The extracellular gel tends to clump once fixed and may stay as one piece when mounting. Care

should be taken to quickly but efficiently resuspend the gel.

15. Allow the agarose to polymerize.

16. Fill the chamber with PBS prior to introducing the sample for imaging.

3.7 Preparation of a Whole Mount of a Mosquito (Anopheles gambiae)

Adult insects are very large objects that can be up to 5 cm long. Moreover, they have an exoskeleton made of chitin which is hydrophobic and autofluorescent. This characteristic allows imaging without labeling by simply using the chitin autofluorescence to image the insect surface. This protocol applies to every type of adult insects as well as for various type of plankton.

Equipment and reagents:

− An adult Anopheles gambiae

− 1.5 % Low Melting Point (LMP) agarose in water or PBS

− 1 ml syringe (BD Biosciences)

− Ethanol (70 %)

− Glycerol (50 %)

− Sucrose

− Heating block

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Fig. 19 Mounting a complete adult Anopheles gambiae.

(A) The insect is paralyzed. (B) The chitin surface is treated using 70 % ethanol. (C) The insect is positioned within the melted agarose. (D) The mounted insect is ready for imaging.

Method:

− Choose an adult Anopheles gambiae and immobilize it by cold treatment. Melt 1.5 % LMP agarose, aliquot 0.5 ml into a 1.5 ml Eppendorf tube. Invert the tube to mix and allow agarose to cool to 40º C.

− To avoid bubble formation on the insect surface that will affect imaging, the insect must be treated either with ethanol 70 % (animal death) or using 50 % glycerol or 1 M sucrose to cover the hydrophobic chitin surface prior to embedding (Fig. 19/B).

− The syringe is prepared as previously described and filled with molten low melting point agarose (40 °C).

− The insect can then be inserted into the agarose cylinder and aligned using a needle or forceps (Fig. 19/C).

− The insect can then be imaged (Fig. 19/D).

This technique can be applied to any insect or similar type of organism possessing an exoskeleton.

Depending on the animal part to be observed the insect can be aligned differently or dissected prior to embedding (head, wings, guts, salivary glands…).

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4 Tips, Troubleshooting and Additional Information

4.1 Tips

Protocols

As a general rule, follow the protocol carefully and try it out well in advance as you may encounter difficulties and missing parts (chemicals…) that may hinder you to use the sample straight away. The protocols presented in this section are not the ultimate solution for every problem. They can be improved and adapted to your need. If you encounter a problem, check the scientific literature and the other protocols presented and see if you can find a tip that you may apply to solve your problem. Be creative.

Safety

Please refer to the safety instructions provided with the Lightsheet Z.1.

Setup Requirements and Maintenance

For optimal performance of the Lightsheet Z.1, please check the Setup Requirements information delivered with the system.

For information about maintenance and cleaning any part of the Lightsheet Z.1, including the optics, please refer to the according CHAPTER in this manual.

Refractive index mismatch

Light is refracted when it crosses the interface between two media of differing refractive indices (RI). Mismatching the refractive index of the objective immersion medium and mounting medium is one of the main causes of image degradation in microscopy. Refractive index mismatch results in stretching/compression of the z-axis. Also, spherical aberration is worsened by axial spreading of the point-spread function (PSF) resulting in reduced axial resolution. This phenomenon is exacerbated with depth and with a high numerical aperture objective, serious problems arise when imaging deeper than 10 µm into an aqueous sample. The mounting medium and the immersion medium should be matched. It is not a major issue in LSFM but this as to be considered when filling the imaging chamber in regard to the sample preparation technique, especially for embedding as gelling agents are used.

4.2 Troubleshooting

Common problems

LSFM is a fluorescence microscopy technique so many troubleshooting guidelines from other microscopy techniques apply here as well. Do not hesitate to ask experts in the field, check the literature, as well as internet resources that may provide you with a more detailed solution to the problem you have encountered. Also check with your Lightsheet Z.1 specialist for FAQs and tips for troubleshooting.

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Sample image is unclear, blurred or has insufficient contrast.

− This can be a simple optical problem: objectives or filters are dirty. Clean them accordingly. You can also check that all the components are well in place and aligned.

− Your light sheet might not be properly aligned. For Lightsheet Z.1, please realign the light sheet using the Light sheet auto-adjust function of the ZEN software.

− You could be imaging through an additional layer of material: glass, plastic... that belong to the mounting support and not the sample itself. Please, check the sample position or move it around to see if another angle solve the problem as a piece of glue, additional agarose or part of the mounting material is having a blurring effect.

− In the case of sample embedding it may occur that the gelling media is of low quality or badly polymerized. This leads to an uneven polymerization of the media that modify the optical path. Try again.

− The light sheet comes from the side and any obstacle modifies its quality. Check the illumination axis for any obstacle (capillary, objects…).

− Check if the medium level in the sample chamber has dropped below the imaging level. The specimen has to be fully immersed for good image quality.

Sample image is partially obscured or unevenly illuminated.

− This can be a simple optical problem: objectives or filters are dirty. Clean them accordingly. You can also check that all the components are well in place and aligned.

− Your light sheet might not be properly aligned. For Lightsheet Z.1, please realign the light sheet using the Light sheet auto-adjust function of the ZEN software.

− The light sheet comes from the side and any obstacle modifies its quality. Check the illumination axis for any obstacle (capillary, objects…).

Sample signal is weak.

− Human eyes have trouble quickly adjusting to the dark and it will be hard to discern a very dim fluorescent specimen immediately after darkening the room. You may want to check your sample using another microscope or a stereomicroscope equipped with a fluorescent lamp.

− It could be an optical problem, e.g. extra filter in the optical path, misaligned illumination or dying laser. You can also increase excitation energy (laser). However, the risk of bleaching and signal saturation will increase.

− In the case of immunofluorescence, you should increase antibody concentration or incubation time but this might in turn increase nonspecific background signal.

− In the case of GFP signal, the expression level might be too low or you have photobleached or damaged the GFP signal during sample preparation (fixation, ethanol treatment, excessive illumination during dissection...).

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High background signal within the sample.

− In the case of immunofluorescence, you should decrease antibody concentration or incubation time but this may decrease the overall signal. You can also use blocking steps during the immunofluorescence (eg. BSA, FCS…). If preparing tissue section, you should increase the stringency of the washing steps.

− The imaging chamber may be dirty as well as the media inside. This contamination affects the quality of the light sheet and scattering occurs.

− You can apply deconvolution to your stacks afterwards.

The sample is moving.

− If you are imaging life samples, it may be simply due to life. The sample is moving so you may want to increase the anesthetic concentration or the agarose concentration to restrain any movement.

− The mounting is unstable. This occurs if the chosen material is not properly maintained (bad tweezers, leaking plunger, badly polymerized agarose…). For embedded sample preparation, you can improve the stability by limiting the amount of agarose emerging out of the syringe or the capillary, as the longer is the agarose tube the more unstable it will be. You can also tighten the plunger by sealing it with nail polish to avoid air leakage that will lead to gliding of the agarose tube.

− The system table might not be a float. Check that it is connected to the pneumatic supply and the air-dampening is active.

− The cables are not properly secured with the cable holders at the system table.

− Other instruments that produce vibrations, not completely dampened by the system table, are in

close proximity (e.g. fridges, centrifuges, etc.).

− The stage is not properly fitted or damaged and prone to vibrations. This includes the sample holder and the imaging chamber support.

Optical aberrations

− As in any optical technique, the LSFM has advantages and disadvantages. Some optical aberrations are more generals and can be dealt with easily. Do not hesitate to ask experts in the field, check the literature as well as Internet resources that may provide a more detailed solution to the problem you have encountered.

− A few optical aberrations are, however, typical for LSFM, as the optical axis are at a 90 degrees angles. Lines and stripes occur as any objects blocking the light sheet will reduce the light intensity leading to a discrepancy along the illuminated plane. This is often a problem with big samples, highly scattering samples or sample which have absorbing structures at the surface of the specimen volume. Rotating the sample to give a better path for the light sheet should be considered first. Dual Side illumination and/or Pivot scanning of the light sheet can often eliminate these effects (available in the Lightsheet Z.1) Second, the sample could be orientated differently during mounting or partially dissected to limit obstacles. The concentration of the objects can also be a problem especially with samples with optical properties (beads, tubes, glass capillaries…). For example, if you image large number of cell clusters, you may have a few of them in the light sheet path. By reducing the amount of objects, you will automatically reduce the lines and stripes.

− The use of image processing methods such as deconvolution may help to get rid of those aberrations.

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4.3 Suggested Additional Sources of Information

Chemicals

• Agarose:

− Molecular biology grade, for routine use, SIGMA, Ref. A-9539

− Type VII, low gelling temperature, SIGMA, Ref. A-4018-50G

− "Agarose Low Melt" (no 6351.1) from Carl Roth http://www.carlroth.com (from the US please

contact Brunschwig Chemie B.V., Amsterdam, NL (e-Mail: brunschwig@brunschwig.nl)

• Gelrite Gellan Gum, SIGMA, Ref. G-1910

• Glycerine anhydrous, AppliChem, Ref. A3552,1000

• Nail Polish, any cosmetic shop near you

• PBS, local supplier

• Distilled water, local supplier

• Ethanol, local supplier

• Companies:

− Sigma- Aldrich (http://www.sigmaaldrich.com/)

− Applichem (http://www.applichem.de/)

− MP BIomedicals (http://www.mpbio.com/)

− Merck KGa (http://www.merck.de/)

Materials

• Capillaries

− 100 µl, color code Blue, Brand GmbH, Ref. 7087 45

− 200 µl, color code Red, Brand GmbH, Ref. 7087 57

• Companies:

− Brand GmbH (http://www.brand.de/en/home/)

− SpectraGlass (http://www.spectraglass.com/)

− Harvard Apparatus (http://www.harvardapparatus.com/)

• Syringes

− 1 ml, BD Plastipak, BD Biosciences, Ref.300013

− 0.5 ml, BD Microfine Insulin U100 Syringe, 29 g, Ref. PLA257L

− 0.3 ml, BD Microfine Insulin U100 Syringe, 30 g,Ref. PLA470U

− Braun Omnifix F Solo 1 ml Syringe (PZN 0569881, Ref. 61706)

− Terumo 1 ml Syringe (Ref. BS-01T)

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• Pipettes

− 2 ml serological pipette, FALCON, BD Labware, Ref.35 7507

− 1 ml serological pipette, FALCON, BD Labware, Ref.35 7521

Equipments

For the following, please refer to your local lab supply companies

• Heating blocks

• Stereomicroscope

• Scalpels

• Tweezers

• Dissection Needles

• Watch glass

• Sonicator

4.4 References and Further Reading

Buytaert, J.A.N. et al., 2011. The OPFOS Microscopy Family: High-Resolution Optical Sectioning of Biomedical Specimens. Anatomy research international, 2012.

Capoulade, J. et al., 2011. Quantitative fluorescence imaging of protein diffusion and interaction in living cells. Nature biotechnology, 29(9), pp.835–839.

Dodt, H.U. et al., 2007. Ultramicroscopy: three-dimensional visualization of neuronal networks in the whole mouse brain. Nature methods, 4(4), pp.331–336.

Ejsmont, R.K. et al., 2009. A toolkit for high-throughput, cross-species gene engineering in Drosophila. Nature methods.

Engelbrecht, C.J. et al., 2007. Three-dimensional laser microsurgery in light-sheet based microscopy (SPIM). Optics Express, 15(10), pp.6420–6430.

Engelbrecht, C.J., Voigt, F. & Helmchen, F., 2010. Miniaturized selective plane illumination microscopy for high-contrast in vivo fluorescence imaging. Optics letters, 35(9), pp.1413–1415.

Fahrbach, F.O. & Rohrbach, A., 2010. A line scanned light-sheet microscope with phase shaped selfreconstructing beams. Optics Express, 18(23), pp.24229–24244.

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Fuchs, E. et al., 2002. Thin laser light sheet microscope for microbial oceanography. Optics Express, 10(2), p.145.

Hama H, Kurokawa H, Kawano H, Ando R, Shimogori T, Noda H, Fukami K, Sakaue-Sawano A, Miyawaki A. 2011. Scale: a chemical approach for fluorescence imaging and reconstruction of transparent mouse brain. Nat Neurosci. 14(11):1481-8.

Hofman, R. et al., 2008. Morphology and function of Bast’s valve: additional insight in its functioning using 3D-reconstruction. European Archives of Oto-Rhino-Laryngology, 265(2), pp.153–157.

Hofman, R., Segenhout, J. & Wit, H., 2009. Three-dimensional reconstruction of the guinea pig inner ear, comparison of OPFOS and light microscopy, applications of 3D reconstruction. Journal of microscopy, 233(2), pp.251–257.

Holekamp, T.F., Turaga, D. & Holy, T.E., 2008. Fast three-dimensional fluorescence imaging of activity in neural populations by objective-coupled planar illumination microscopy. Neuron, 57(5), pp.661–672.

Huber, D., Keller, M. & Robert, D., 2001. 3D light scanning macrography. Journal of microscopy, 203(2), pp.208–213.

Huisken, J. et al., 2004. Optical sectioning deep inside live embryos by selective plane illumination microscopy. Science, 305(5686), pp.1007–1009.

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CHAPTER 2 - SAMPLE PREPARATION Lightsheet Z.1 Tips, Troubleshooting and Additional Information Carl Zeiss

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CHAPTER 2 - SAMPLE PREPARATION Carl Zeiss Index Lightsheet Z.1

5 Index

Antifading Agents ......................................... 26

FEP Tubing .................................................... 17

Fixation and Fixatives ..................................... 25

LSFM

Sample Mounting ..................................... 6

Materials and Equipments

Sample Chambers .................................. 18

Molding and Mounting Supports ............ 20

Sample Holder ....................................... 22

Gels and Polymers .................................. 23

Hydrogel Preparation .............................. 24

Sample

Holding ................................................... 8

Sample Preparation

Preparation of Fluorescent Beads .............28

Preparation of a Medaka Fish Embryo ......29

Preparation of a Fly Pupa .........................31

Preparation of a Plant Root ......................32

Samples

Embedded ..............................................10

Hanging .................................................14

Enclosed .................................................15

Cleaning, Labelling, and Storing ..............26

Stains and Staining .........................................25

44 000000-1790-528 02/2013

Zeiss Lightsheet Z.1 Operating Manual

Specifications and Main Features

Frequently Asked Questions

User Manual

Contents of this Manual

CHAPTER 1 - HARDWARE Lightsheet Z.1 Content Carl Zeiss

1 Maintenance and Cleaning

1.1 Maintenance of the Liquid Cooling System

2 ErgoDrive Operating Panel

3 User Interfaces

3.1 Installation and Deinstallation of the Detection Modules

3.2 Adjustment – Detector Recognition

3.2.1 Adjustment – Automatic Detector Alignment

3.2.2 Adjustment – Manual Detector Alignment

3.2.3 Adjust the Grating Focus for the Automatic or Manual Detector Alignment Tool

3.2.4 Cable Connections for the Detection Module "PCO.Edge"

3.2.5 Cable Connections for the Detection Module "Standard"

3.3 Assembly of the Sample Chamber

3.3.1 Assembly of the Sample Chamber Windows

3.3.2 Assembly of the Sample Chamber Body and the Sample Chamber Dove Tail Slide

3.3.3 Insertion of the Drain Connector, Luer-Lock Connectors and Blind Plugs

3.3.4 Insertion of Accessories for Incubation

3.4 Removing and Inserting the Sample Chamber

3.5 Assembly of the Sample Holder

3.5.1 Assembly of the Sample Holder for Capillary

3.5.2 Assembly of Sample Holder for Syringes

3.6 Inserting and Removing the Sample Holder

3.7 Installation of the Incubation Modules

3.7.1 Heating Components

3.7.3 Heating Device Humidity S1

3.7.4 Registration of the incubation modules

3.8 Switch Incubation ON and OFF

3.9 Illumination and Detection Optics

3.9.1 Removing and Inserting the Illumination Optics Unit

3.9.2 Removing and Inserting the Detection Optics Unit

3.10 Removing and Inserting the Reflector Turret for Emission Selection

3.11 Removing and Inserting the Reflector Turret for Laser Blocking Filter

4 Index

CHAPTER 2 - SAMPLE PREPARATION Lightsheet Z.1 Content Carl Zeiss

1 Introduction

2 Sample Mounting for LSFM

2.1 The perfect Sample for LSFM

2.2 Holding the Sample

2.2.1 Embedded Samples

2.2.2 Hanging Samples

2.2.3 Enclosed Samples

2.2.4 FEP Tubing

2.3 Materials and Equipment

2.3.1 Sample Chambers

2.3.2 Molding and Mounting Supports

2.3.3 Sample Holder

2.3.4 Gels and Polymers

2.3.5 Hydrogel Preparation

2.4 Fixation and Fixatives

2.5 Stains and Staining

2.5.1 Choosing a Fluorescent Label

2.6 Antifading Agents

2.7 Cleaning, Labelling and Storing Samples

3 Specific Examples of Sample Preparation

3.1 Preparation of Fluorescent Beads

3.2 Preparation of a Medaka Fish Embryo (Oryza latypes)

3.3 Preparation of a Fly Pupa (Drosophila melanogaster)

3.4 Preparation of a Plant Root (Arabidopsis thaliana)

3.5 Imaging Cell Cysts in an Extracellular Matrix Gel

3.6 Immunostaining and Preparation of MDCK Cell Cysts

3.7 Preparation of a Whole Mount of a Mosquito (Anopheles gambiae)

4 Tips, Troubleshooting and Additional Information

4.1 Tips

4.2 Troubleshooting

4.3 Suggested Additional Sources of Information

4.4 References and Further Reading

5 Index