Here we present a protocol to isolate nuclei from the brains of the short-lived vertebrate model Nothobranchius furzeri for downstream applications such as single-nucleus RNA sequencing or single-nucleus assay for transposase-accessible chromatin with sequencing (ATAC-seq).
Studying brain aging at single-cell resolution in vertebrate systems remains challenging due to cost, time, and technical constraints. Here, we demonstrate a protocol to generate single-nucleus RNA sequencing (snRNA-seq) libraries from the brains of the naturally short-lived vertebrate African turquoise killifish Nothobranchius furzeri. The African turquoise killifish has a lifespan of 4-6 months and can be housed in a cost-effective manner, thus reducing cost and time barriers to study vertebrate brain aging. However, tailored protocols are needed to isolate nuclei of sufficient quality for downstream single-cell experiments from the brain of young and aged fish. Here, we demonstrate an empirically optimized protocol for the isolation of high-quality nuclei from the brain of adult African turquoise killifish, a critical step in the generation of high-quality single nuclei omic libraries. Furthermore, we show that the steps to reduce contaminating background RNA are important to clearly distinguish cell types. In summary, this protocol demonstrates the feasibility of studying brain aging in non-traditional vertebrate model organisms.
Understanding the mechanisms of vertebrate brain aging is critical to addressing age-related neurodegenerative diseases such as Alzheimer's and dementia1. The African turquoise killifish (Nothobranchus furzeri) is the shortest-lived vertebrate that can be bred in captivity, and due to its short lifespan and age-associated cognitive impairment, it is an excellent brain aging model2,3,4,5. Recently, the advent of single-cell "omics" technologies, such as single nuclei RNA-seq (snRNA-seq) and single nuclei assay for transposase-accessible chromatin with sequencing (snATAC-seq), have allowed researchers to interrogate the aging brain at an unprecedented resolution6,7,8. These methods rely on nuclei isolation, since the recovery of brain cells such as neurons is often too challenging to isolate6,7,8,9,10. However, most published nuclei isolation protocols are optimized for mammalian model organisms11,12,13,14,15. Thus, as there is currently an unmet need for isolating brain nuclei in the killifish as an up-and-coming new model organism in the field of aging research2, the goal of this protocol is to establish a method for isolating high-quality nuclei from frozen brain killifish tissue.
Here, a streamlined and robust workflow is established that uses commonly available materials to isolate high-quality nuclei from killifish brains. This protocol was modified from a 10x Genomics protocol for mouse brains to accommodate the lower myelin content brains of African turquoise killifish, the fragility of frozen tissue, and the need to reduce ambient debris content for sequencing-related applications16. Indeed, previously optimized protocols for mammalian brain tissue17,18 lead to poor nuclei quality (i.e., overlysis) and/or high debris content when used on frozen killifish brains, making them unsuitable for use with snRNA-seq according to recommendations for single nuclei RNA-seq using microfluidics (Supplementary Figure 1).
In addition to nuclei isolation, we demonstrate how to assess nuclei quality and yield by microscopy and flow cytometry. This article provides examples of both optimal and suboptimal results and discusses troubleshooting. This protocol was designed and optimized for frozen killifish brains but can also be used without major modifications on freshly dissected killifish samples. Killifish brain nuclei isolated using this method have been optimized for use in single nucleus RNA-seq (snRNA-seq) as a downstream application, but should also be amenable for use in snATAC-seq and bulk ATAC-seq.
Animal care and animal experimentation were performed in accordance with the University of Southern California IACUC under approved protocols #21215. For any work using this protocol, it is necessary to obtain approval from the institution's IACUC prior to starting any research work on vertebrate animals.
NOTE: A complete run through the protocol starting from flash-frozen brain tissue (starting from step 3) should take ~2.5 h for six samples (Figure 1).
1. Dissect killifish brains
2. Prepare fresh buffers
3. Isolate nuclei
4. Assess nuclei quality by microscopy
5. Quantify singlet nuclei, debris, and multiplet proportion by flow cytometry
NOTE: The specific terminology and interface of the flow cytometer software may differ based on the brand of the machine, but these steps should be easily adapted to other systems if required.
The killifish brain nuclei isolation protocol described here is optimized specifically for the killifish and is summarized in Figure 1. In addition to nuclei extraction, the protocol details different methods for assessing the quality and quantity of isolated nuclei. Figure 2 shows examples of both healthy (Figure 2A) and unhealthy (Figure 2B) nuclei as assessed by light microscopy. Healthy nuclei suitable for downstream analysis (Figure 2A) present as singlet nuclei with intact membranes. As shown in Figure 2B, the most common failure mode of this protocol (and other nuclei isolation protocols) is the generation of overlysed nuclei, which are characterized by damaged nuclear membranes. This often leads to nuclei clumping and can contribute to background signals in downstream applications due to the leakage of nucleic acids from the ruptured nuclei. If excessive clumping is observed, we recommend being gentler during the dounce steps and/or reducing the incubation times in lysis buffer.
This protocol uses flow cytometry to quantify the number of isolated nuclei and determine the relative proportion of multiplet nuclei in a sample. Additionally, flow cytometry can be used to assess the relative content of contaminating debris, often ruptured nuclei fragments and other cellular debris that can detrimentally affect downstream assays in the nuclei sample. Representative flow data can be seen in Figure 3. A high-quality nuclei isolation procedure will produce: 1) a small debris fraction, and 2) a high fraction of singlet versus multiplet nuclei (> 80% of the nuclei fraction). The use of PI staining, which stains nucleic acids, allows singlet nuclei to be segregated from debris and multiplet nuclei. Figure 3A is an example of a prep contaminated by debris, whereas Figure 3B is an example of a high-quality experiment.
Figure 1: Killifish brain nuclei isolation workflow. Experimental workflow for nuclei isolation from frozen or fresh killifish brains. Once isolated and assessed, nuclei can be used for various downstream "omics" analyses. Please click here to view a larger version of this figure.
Figure 2: Assessment of nuclei quality by microscopy. Nuclei were mixed 1:1 in a solution of 0.4% Trypan Blue and visualized on an Echo Revolve microscope by brightfield imaging at 60x. (A) An example of a high-quality nucleus. The nucleus is present as a singlet and the nuclear membrane is intact. (B) An example of poor-quality nuclei. The nuclear membranes are damaged, and nuclei are clumping into a doublet, likely due to leakage of sticky DNA, which can be seen leaking from the top of the topmost nucleus. Please click here to view a larger version of this figure.
Figure 3: Nuclei quantification and purity assessment by flow cytometry. Nuclei samples were stained with a 1:100 dilution of PI and run on a flow cytometer. Nuclei are present in singlet and multiplet forms in the upper right quadrant of (A,B) with singlets as the lowest cloud of events followed by doublets, triplets, etc. in ascending order. Debris is marked by the events in the leftmost two quadrants. (A) An example of a low-quality nuclei prep, where the debris removal step was omitted and debris makes up >40% of events. (B) An example of a high-quality nuclei sample, with the included debris removal step. In this example, debris makes up <10% of all events registered by the flow cytometer. Please click here to view a larger version of this figure.
Sample type (2 Brains) | Average Yield (4 independent preps) |
5-week Male Brains | 3.59 ± 1.76 x 105 |
10-week Male Brains | 6.41 ± 1.33 x 105 |
15-week Male Brains | 14.59 ± 2.05 x 105 |
5-week Female Brains | 2.54 ± 0.75 x 105 |
10-week Female Brains | 4.66 ± 1.29 x 105 |
15-week Female Brains | 7.95 ± 3.51 x 105 |
Table 1: Average expected yields from two brains of African turquoise killifish across sex and age. Average yields expressed as 105 nuclei ± standard error of the mean over four independent nuclei preparations in each category.
Supplementary Figure 1: Comparison of nuclei isolation quality using standard protocols and our optimized protocol on frozen killifish brains. (A) Nuclei quality assessment by microscopy. Nuclei were mixed 1:1 in a solution of 0.4% Trypan Blue and visualized on a microscope by brightfield imaging at 60x. (B) Nuclei and debris load quantification by flow cytometry. Nuclei samples were stained with a 1:100 dilution of PI and run on a flow cytometer. (C) Summary of quality assessment metrics by microscopy and flow cytometry with the different benchmarked protocols. Please click here to download this File.
Supplementary Figure 2: Schematic representation of the debris removal steps. (A) Scheme of debris removal layering pre-centrifugation. (B) Scheme of supernatant removal after centrifugation. Please click here to download this File.
The protocol presented here can be used to reproducibly generate high-quality nuclei from killifish brains. This protocol had to be specifically designed for the killifish brain as typical mammalian-based brain nuclei isolation protocols applied to killifish brains consistently resulted in poor nuclei quality in our hands. We suspect that this is due to the lower relative myelin content of the killifish brain compared to their mammalian counterparts, which would lyse and clump in response to the harsh conditions required for mammalian brain cell lysis. This protocol is an advancement in the aging and killifish fields as it facilitates the exploration of brain aging at the single-cell level in a cost and time-effective model of vertebrate brain aging.
This protocol is robust to fresh or frozen samples, though one must consider the downstream applications when using fresh or frozen tissue. Frozen tissue is often convenient as it can be collected and stored for months while samples are collected. Such samples can be safely used for applications such as snRNA-seq. However, freezing samples may disrupt the nuclear structure and thus the ability to accurately measure the chromatin landscape by ATAC-seq21. Thus, for downstream applications such as bulk ATAC-seq or snATAC-seq, it is recommended to use freshly dissected brains instead of frozen brains. In addition, because all steps after brain homogenization can be performed in parallel, this protocol is amenable to running multiple samples in a reasonable timeframe, thus limiting RNA degradation caused by prolonged incubation on ice.
Furthermore, it is imperative to prepare buffers fresh (within hours) of performing nuclei extraction. We found that detergents as well as BSA must be added to buffers immediately prior to beginning the protocol. Buffers containing only salts (PBS, NaCl, etc.) may be made as concentrates, filter sterilized (0.22 µm), and stored indefinitely at room temperature. BSA stock solutions may be prepared, sterilized, and stored at 4 °C within days of the nuclei extraction (if prepared from a powder) but should always be added to the buffers used in this protocol immediately before undertaking the protocol. However, we recommend preparing BSA solutions on the day of the protocol. If using premade BSA solutions from a third party, it is advised to use fresh, unopened bottles. Using fresh BSA generally leads to lower debris content in nuclei preps.
Whether fresh or frozen samples are used as input, it is important to assess nuclei quality following nuclei isolation. Though this protocol is specifically designed to avoid overlysis, this is the most common cause of nuclei quality loss. Overlysis may result from too much time spent in the lysis buffer, overly rough handling of the nuclei such as excessive pipetting with a standard bore pipette tip, or an excessive amount of time spent between nuclei isolation and downstream applications (>1 h). Overlysed nuclei will often have damaged nuclear peripheries, which leak DNA and cause clumping (Figure 2B). This will lead to an increased number of multiplets and contribute background nucleic acids that will interfere with downstream applications, especially snRNA-seq. Both qualities can be assessed by microscopy following nuclei isolation. If excessive nuclear clumping is observed, we recommend trying to shorten the lysis step incubation to reduce the chance of overlysis. As an alternative method for increasing nuclei singlets, fluorescence-assisted cell sorting (FACS) may be used to enrich for singlets downstream of this protocol. However, we note that, when working with already fragile nuclei from frozen tissue, the shear stress occurring during sorting may lead to increased nuclear rupture and thus increased ambient RNA/DNA. In addition, we note that the time required to run a FACS yield sort for nuclei when processing multiple samples would require all nuclei samples to remain on ice for hours, while other samples are being sorted. Thus, increased wait times when processing multiple samples in parallel with the FACS approach could also likely lead to overall reduced nuclei quality and increase the risk of RNA degradation. Thus, if FACS is desired for reduced doublet rate, we recommend that debris content should be checked again after the yield sort and that possible reduced RNA quality be taken into account for single cell RNA-seq applications as a potential caveat.
An accurate estimate of nuclei counts and singlet proportion is essential for nearly all downstream "omics" applications and is of the utmost importance. Due to the ability to easily gate and count nuclei by size, flow cytometry is the most accurate method of counting nuclei that we have assessed. Alternatively, one may quantify nuclei using cell counters such as Invitrogen's Countess 2 FL Automated Cell Counter or the DeNovix CellDrop Automated Cell Counter. To note, Invitrogen's Countess 2 FL Automated Cell Counter, and to a much lesser extent the DeNovix, tend to overestimate nuclei counts by counting debris as nuclei, which means that manual size gating may be required. Furthermore, the flow cytometer allows one to easily assess the purity of the nuclei. One can discern the relative proportion of singlet versus multiplet nuclei in a quantitative manner that is difficult by microscopy. This is vital for snRNA-seq and snATAC-seq, since those protocols will suffer from a surplus of multiplet samples, which must be excluded from downstream analyses. In addition to multiplets, the relative proportion of "debris" (fragmented nuclei, cellular debris) can be quantified by flow cytometry and must be relatively low, since this material often contains contaminating nucleic acids that can contribute a background signal and corrupt single nucleus "omics" data.
As with previously described nuclei isolation protocols, the proportions of cell types in the original tissue may not be recapitulated faithfully in the nuclei prep19, and thus should be interpreted with caution. To note, like all teleosts, African turquoise killifish have nucleated erythrocytes, which are also expected to be represented in the nuclei prep. These nuclei can be identified in snRNA-seq and snATAC-seq datasets by the higher expression/accessibility of hemoglobin genes and may be excluded computationally if desired.
Some panels were generated with BioRender.com. This work in our laboratory was supported by the NIA T32 AG052374 Postdoctoral Training Grant to B.T., a grant from the Simons Foundation as part of the Simons Collaboration for Plasticity in the Aging Brain, a pilot grant from the Navigage Foundation, and a Hanson-Thorell Family award to B.A.B.
Name | Company | Catalog Number | Comments |
0.4% Trypan Blue solution | Gibco | 15250061 | |
10x PBS | Bioland | PBS01-03 | |
15 mL Conicle Centrifuge Tube | VWR | 89039-664 | |
2 mL Tissue Grinder | Kimble | 885300-0002 | Dounce Homogenizer |
5 mL Polystyrene Round-Bottom Tube | Falcon | 352054 | |
5M Sodium Chloride, Molecular Biology Grade | Promega | V4221 | |
autoMACS Rinsing Solution | Miltenyi Biotec | 130-091-222 | This corresponds to 1x PBS pH 7.2, 2 mM EDTA (used for flow cytometry) |
Debris Removal Solution | Miltenyi Biotec | 130-109-398 | |
DNA LoBInd Tube | Eppendorf | 22431021 | |
Echo Revolve microscope (fitted with 60x objective) | Echo | NA | No catalog number; Used to visually inspect nuclei. |
Falcon Round-Bottom Polystyrene Tubes, 5 mL | Falcon | 352052 | FACS tube |
FLOWMI Cell Strainers, 40 μM | SP Bel-Art | 136800040 | Referred to as on-tip filters in Protocol |
Hydrochloric Acid | Sigma-Alrdich | 258146-500 mL | Used to lower TRIS pH from 8.0 to 7.4 |
Leica EZ4 dissecting scope | Leica | NA | No catalog number |
MACS BSA stock solution | Miltenyi Biotec | 130091376 | |
MACS SmartStrainers (70 μM) | Miltenyi Biotec | 130-110-916 | |
MACSQuant Analyzer 10 Flow Cytometer | Miltenyi Biotec | NA | No catalog number |
Magnesium Chloride, Hexahydrate, Molecular Biology Grade (Powder) | Millipore | 442611 | |
Megafuge 16R Centrifuge | ThermoScientific | 75003629 | |
Micro Cover Glass | VWR | 48393081 | |
Micro Slides Superfrost Plus | VWR | 48311-703 | |
Nonidet P-40 Substitute | Roche | (Roche) 11332473001/ Catalog: 983739 P Code: -102368106 (Sigma Aldrich) | |
NP-40 Surfact-Amps Detergent Solution | ThermoFisher | 85124 | |
Nuclease-Free HyPure Molecular Biology Grade Water | HyClone | SH30538.02 | |
NxGen RNase Inhibitor (50,000 U) | Lucigen | 30281-2 | |
Propidium Iodide solution | MBL | FP00010020 | |
PureBlu DAPI Nuclear Staining Dye | Biorad | 1351303 | |
TipOne RPT Pipette Tips (Ultra low retention, filtered) in 10 µL, 20 µL, 200 µL, and 1000 µL sizes | USA Scientific | #1181-3810; #1180-1810; #1180-8810; #1182-1830 | |
Tricaine-S (MS 222) | Syndel | Tricaine10G | Syndel is an FDA-approved provider for pharmaceutical grade Tricaine |
TRIS, 1 M, pH 8.0 | VWR | E199-500 mL | |
Wide Bore Pipet Tips | Axygen | T-1005-WB-C |
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