Drosophila Late Pupa Indirect Flight Muscle (IFM) Dissection: A Method for High-Throughput Tissue Collection

Abstract

Source: Kao, S. Y., et al. Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches. J. Vis. Exp. (2019).

The Drosophila flight muscles are used as a model system for muscle development and physiology. This video describes the flight musculature of adults and highlights a protocol to dissect developing indirect flight muscles that are suitable for RNA sequencing from pupae.

Protocol

This protocol is an excerpt from Kao et al., Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches, J. Vis. Exp. (2019).

1. IFM Dissection After 48 h APF

1. Assemble necessary equipment including two #5 biology grade forceps, fine scissors, standard glass microscope slides, double-stick tape, pipette, pipette tips, dry ice, and (for RNA applications) isolation reagent (see Table of Materials). Chill the 1x PBS and microcentrifuge tubes on ice.
2. Using a lightly wetted paintbrush, transfer the staged pupae to a strip of double-sided sticky tape mounted on a microscope slide (Figure 1A). Place the pupae in a line, oriented in the same orientation (ventral down and anterior towards the bottom of the slide).
   NOTE: Be careful not to use too much water on the paintbrush or filter, or the pupae will not stick well. If pupae do not stick, dry them by first transferring to a dry filter or tissue paper. Mount as many pupae as can be dissected within a 30 min time window, ideally ~10 pupae.
3. Remove the pupa from the pupal case. Use forceps to tease apart and open the pupal case above the anterior spiracles (Figure 1B).
4. Gently slide a pair of forceps dorsally towards the posterior, cutting the pupal case as the forceps move (Figure 1B'). Be careful not to rupture the underlying pupa. Liberate the pupa from the opened case and immediately transfer it to a drop of 1x PBS on a second microscope slide (Figure 1B",C).
5. Repeat steps 1.3 and 1.4 for all pupae in the line, then set the double-stick tape slide aside.
6. Using the fine scissors, cut the abdomen of the pupa away from the thorax and push it into a separate pile (Figure 1D,D'). Repeat for the remaining pupae.
   NOTE: Begin timing the length of dissection with step 1.6, as soon as pupal integrity is disrupted. Dissect as many flies as possible in 20–30 min to prevent cell death and associated transcriptomic and proteomic changes. When dissecting 1 d adults or >90 h pupae, it is often convenient for later steps to additionally remove the head with the fine scissors.
7. Using a tissue paper, remove the majority of the 1x PBS (generally cloudy with suspended fat) as well as the pile of abdomens (Figure 1E). Add a drop of fresh, chilled 1x PBS to the remaining thoraxes.
8. Use the scissors to cut the thorax in half (Figure 1F,F') by cutting from the head down the longitudinal body axis in a single motion. Alternately, if the head has been removed, first insert the scissors where the head was attached and cut the top half of the thorax longitudinally between the IFMs. Then, cut the ventral side of the thorax with a second cut in the same orientation.
9. Repeat steps 1.7 and 1.8 for all pupae to be dissected, generating a pile of thorax hemisections near the center of the slide. Ensure there is enough chilled 1x PBS on the slide so that the hemisections do not dry out.
   NOTE: After 48 h APF, IFMs are large enough to be visible under a standard dissecting microscope to the trained eye. At this point in the protocol, muscles with a fluorescent label can be moved to a fluorescent dissecting scope to aid in IFM identification or for training purposes, but this is not necessary.
10. Dissect the IFMs out of the thorax. Isolate one of the hemisections using the #5 forceps (Figure 1G,H). Gently insert the tips of one forceps above and below the middle of the IFMs (Figure 1G',H'). While holding the first forceps still, use fine scissors to cut one end of the IFM away from the cuticle and tendons. Then, cut the other end of the IFM free from the cuticle (Figure 1G'',H'').
    NOTE: Depending on the orientation of the thorax after the first IFM cut, it is useful to rotate the thorax 180° so that the second IFM cut is easier to perform.
11. Remove the IFM bundle from the thorax with forceps (Figure 1G''',H'''), transferring it to the edge of the PBS bubble to use water tension to hold it in place (Figure 1I). Push the carcass to the opposite side of the slide. Repeat for the remaining thorax hemisections, generating a collection of dissected IFMs.
    NOTE: If the IFMs do not stay in a neat pile, remove some of the 1x PBS with a tissue. Be careful not to let all of the PBS evaporate, and ensure that the dissected IFMs and hemithoraxes remain covered by buffer.
12. After dissecting all IFMs, quickly perform a quality control on the dissected muscle. Using #5 forceps, remove any jump muscle or cuticle fragments that may have found their way into the sample (Figure 1J-K'').
    NOTE: Jump muscle appears different from IFM. If dissecting Mef2-Gal4 labeled muscle under fluorescence, jump muscle has a weaker fluorescence and a different shape and texture. Under normal light, it appears nearly translucent while the IFMs are an opaque, milky yellow (Figure 1J-J'',K).
13. Using water tension, capture (but do not squish) the dissected IFMs between a pair of forceps (Figure 1L). Transfer the IFMs to a 1.5 mL microcentrifuge tube pre-filled with 250 μL of chilled 1x PBS (Figure 1M). Proceed immediately with section 2.
    NOTE: When forceps tips are brought into proximity of each other and lifted out of a buffer solution, water tension causes a bubble of buffer to be captured between the forceps tips. If IFMs are also present in this bubble, they can be lifted out of the solution and easily transferred to another buffer-filled receptacle. It is important to squeeze the forceps to bring the tips near one another without touching each other, to avoid macerating the tissue captured in the buffer bubble.

2. Pellet and Preserve the IFM Sample

1. Pellet the IFMs by centrifuging the 1.5 mL microcentrifuge tube for 3–5 min at 2,000 x g in a table-top centrifuge (Figure 2A,B).
2. Remove the buffer using a pipette tip (Figure 2C).
3. For RNA applications, resuspend the IFM pellet in 50–100 μL of the desired RNA isolation buffer (see Table of Materials, Figure 2D). Otherwise, proceed to step 2.4.
   NOTE: IFMs can be dry-frozen after step 2.2 for mass spectrometry preparations or isolation of RNA with commercial kits (see representative results). For RNA applications, better results are obtained by immediately resuspending and freezing the IFM pellet in isolation buffer.
4. Freeze sample on dry ice or snap freeze in liquid nitrogen (Figure 2E). Store at -80 °C until ready for subsequent steps in sample preparation for downstream analysis.
   NOTE: After cryopreservation, samples can be stored for several months before processing for downstream investigation.

Results

Figure 1: Dissection of IFMs after 48 h APF. (A) Aligning of pupae on double-stick tape. (B) Removal of pupae from the pupal case by opening anteriorly, cutting the case dorsally (B'), and lifting out the pupa (B'...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
60 mm culture dishes	Sigma-Aldrich	Z643084-600EA	Greiner dishes, 60 mm x 15 mM, vented
Cell phone camera, Samsung Galaxy S9	Samsung	SM-G960F/DS	used for photos not taken under a microscope
Double stick tape	Scotch/3M	3M ID 70005108587	Double-sided tape, available at most office supply handlers
Dumont #5 Forceps	Fine Science Tools	11252-20	Inox straight tip 11 cm forceps, Biology grade with 0.05 mm x 0.02 mm tip
fluorescent dissecting microscope camera, Leica DFC310 FX camera	Leica		www.leica-microsystems.com
Fluorescent dissecting microscope software, Leica Application Suite (LAS) version 4.0.0	Leica		www.leica-microsystems.com
Fluorescent dissecting microscope, Leica M165 FC	Leica		www.leica-microsystems.com
Fly: Bru1[M2]			Fly stock; This paper
Fly: Bru1[M3]			Fly stock; This paper
Fly: Mef2-GAL4	Bloomington Stock Center	BDSC:27390	Fly stock
Fly: salm[1]	Bloomington Stock Center	3274	Fly stock
Fly: salm[FRT]			Fly stock; see Spletter et al., Elife, 2018
Fly: UAS-Bru1IR	Vienna Drosophila Research Center	GD41568	Fly stock, RNAi hairpin
Fly: UAS-GFP::Gma	Bloomington Stock Center	BDSC:31776	Fly stock
Fly: UAS-mCD8a::GFP	Bloomington Stock Center	BDSC:5130	Fly stock
Fly: w[1118]	Bloomington Stock Center	3605	Fly stock
Fly: weeP26			Fly stock; see Clyne et al., Genetics, 2003
GFP detection reagent, GFP-Booster	ChromoTek	gba488-100
Glycogen	Invitrogen	10814-010
Image processing software, Photoshop CS6	Adobe		www.adobe.com
Isopropanol	Sigma-Aldrich	I9516-25ML	2-propanol
Method 1 (RNA isolation): TRIzol	Life Technologies	15596018	Guanidinium isothiocyanate and phenol monophasic solution
Method 2 (RNA isolation): Method 1 + TURBO DNA-free Kit	Invitrogen	AM1907	TRIzol isolation followed by treatment with a kit to remove DNA
Method 3 (RNA isolation): Direct-zol RNA Miniprep Plus Kit	Zymo Research	R2070S	RNA isolation in TRIzol, but over commercial columns instead of using phase separation. <br/>Recommended DNase treatment performed with Monarch Dnase I in Monarch DNase I Reaction buffer.
Method 4 (RNA isolation): RNeasy Plus Mini Kit	Qiagen	74134	We used the provided DNase treatment. IFM pellets were homogenized in RTL buffer as suggested for animal tissues.
Method 5 (RNA isolation): ReliaPrep RNA Tissue Miniprep System	Promega	Z6110	We applied the protocol for 'Purification of RNA from Fibrous Tissues'.
Method 6 (RNA isolation): Monarch Total RNA Miniprep Kit	New England Biolabs	T2010G	We applied the protocol for tissues/leukocytes and lysed in 300 &micro;L of RNA Protection Reagent.
Microcentrifuge tubes	Thermo Fisher	AM12400	RNase-free Microfuge Tubes, 1.5 mL
Microscope slides	Thermo Fisher	12342108	Standard slides, uncharged, 1 mm
Paintbrush	Marabu	1910000000	Marabu Fino Round No. 0, or similar brush from any art supply
Paraformaldehyde	Sigma-Aldrich	158127
PBS buffer (1x)	Sigma-Aldrich	P4417	Phosphate buffered saline tablets for 1 L solutions, pH 7.4
PFA PureTip Pipette Tips	Elemental Scientific	ES-7000-0101	Optional substitute for standard pipette tips to reduce sample loss; 100 mL, 0.8 mm orifice
Pipette tips	Sigma-Aldrich	P5161	Universal 200 mL pipette tips
RNA concentration Approach 1 &amp; RNA integrity traces, Bioanalyzer	Agilent Technologies	G2939BA
RNA concentration Approach 2, Nanodrop	Thermo Fisher	ND-2000
RNA concentration Approach 3, Qubit 4 Fluorometer	Invitrogen	Q33238
RNase A	Promega	A7937
RNase-free water, Diethyl pyrocarbonate (DEPC)	Sigma-Aldrich	D5758	DEPC treat water overnight and then autoclave, to remove all RNase.
RT Kit #1: Super Script III Reverse Transcriptase Kit	Invitrogen	18080-044	reverse transcription kit
RT Kit #2: LunaScript	New England Biolabs	E3010S	reverse transcription kit
RT Kit #3: QuantiNova Reverse Transcription Kit	Qiagen	205410	reverse transcription kit
Statistical software: GraphPad Prism	GraphPad Prism		www.graphpad.com
Statistical software: Microscoft Excel	Microsoft		Purchased as part of the bundle: Office Home &amp; Student 2019
Table-top centrifuge	Eppendorf	5405000760	Eppendorf Centrifuge 5425 or equivalent
Tissue/ Kimwipes	Sigma-Aldrich	Z188956	Standard tissue wipes
Transfer pipette	Sigma-Aldrich	Z350796	Plastic pipette
Vannas spring scissors	Fine Science Tools	15000-00	3 mm cutting edge, tip diameter 0.05 mm, length 8 cm
Whatman paper	Sigma-Aldrich	1004-070	Filter paper circles, Grade 4, 70 mm