rainbow capillary experiment

Walking Water Rainbow Science Experiment

Let’s make a walking water rainbow! There’s no better way for little scientists to learn about capillary action and color mixing than by making water walk (yes – walk!) in this colorful rainbow science experiment. This science experiment is a favorite of ours because it’s so easy to set up and the results are almost immediate.

Check out the simple step-by-step below and then gra b 30 more jaw-dropping (but easy prep!) science experiments kids will love from our shop!

Getting Ready

To prep, I gathered our supplies:

• 6 wide-mouth glasses or jars
• Paper towels (use the kind where you can select a size)
• Food dye or liquid water colors (red, yellow, and blue)

I grabbed the six small glasses first .  We’ve had success using wide-mouth drinking cups and canning jars, too.  Even though they all worked, just remember that bigger glasses will need more food coloring.

I ripped off six sheets of paper towel and folded each sheet in thirds, lengthwise.

We were using pretty small glasses, so I cut a few inches off the folded paper towel so it would fit in the glasses.

It’s a good idea to test your paper towel strip to make sure they fit properly in your glasses.  They should be able to go from the bottom of one jar to the next without sticking up in the air too much. The paper towel on the left shows the just-right height.  It’s important to set up this rainbow science experiment for success!

Making a Rainbow

This colorful rainbow science experiment is so simple and quick, it’s perfect for even the youngest little scientists.  My 3 year old, Q, couldn’t wait to get started.

First, I had him line up the glasses and fill the first one with a good squirt of red watercolor , the third with yellow, and the fifth glass with blue.  We left the other glasses empty.

Next, I helped Q add water to the glasses with color until the colored water almost reached the top.

We moved the glasses into a circle and added the paper towels .  Starting with the red, we added one end of the paper towel and then put the other end in the empty glass next to it.

We continued around until the last paper towel was placed into the red glass.

We saw the color wick up the paper towel right away.  This rainbow science experiment doesn’t take long to get going!

After another several minutes, the colored water had almost travelled the whole length of each paper towel.

Five minutes later, the water had traveled all the way up and then down the paper towel and was dripping into the empty glass.

The yellow and red water dripped into the empty cup to make orange!  It made for a good lesson on color mixing.

After another five minutes, we could see the water level had dropped in the red, yellow, and blue glasses and rose in the once empty glasses as the water continued to travel from the more full glasses to the less full glasses.

We grabbed a snack and watched our beautiful rainbow science experiment during the next 20 minutes. The water continued to walk from the primary colored glasses to fill the secondary-colored glasses until all the jars were filled equally.

Not Working?

If you aren’t seeing much movement within a few minutes, it may be that you need to add more water to your colored water glasses.  It really needs to be almost at the top for the water to walk quickly.  So try topping off those glasses and seeing if that gets things moving.

If you see the water moving up the paper towel but it seems like it’s taking forever , it may be the type of paper towel you are using.  You want a paper towel that will really hold a lot of water.  We have used Bounty Select-a-Size and Target’s Up and Up Brand Select-a-Size with success.

It really is worth the extra effort of trying different cups and paper towels to get this activity to work.  And once you have had success, don’t throw out those beautifully-colored paper towels or the colored water!  We gently squeezed out our paper towels and let them dry in a heap on a baking sheet.  We ended up with gorgeous tie-dyed looking paper towels to use for crafts and we used the leftover water as watercolors for painting with later.

The Science Behind It

This rainbow science experiment is as magic as the science behind it.  The colored water travels up the paper towel by a process called capillary action . Capillary action is the ability of a liquid to flow upward, against gravity, in narrow spaces.  This is the same thing that helps water climb from a plant’s roots to the leaves in the tree tops.

Paper towels, and all paper products, are made from fibers found in plants called cellulose .  In this demonstration, the water flowed upwards through the tiny gaps between the cellulose fibers.  The gaps in the towel acted like capillary tubes, pulling the water upwards.

The water is able to defy gravity as it travels upward due to the attractive forces between the water and the cellulose fibers.

The water molecules tend to cling to the cellulose fibers in the paper towel.  This is called adhesion .

The water molecules are also attracted to each other and stick close together, a process called cohesion .  So, as the water slowly moves up the tiny gaps in the paper towel fibers, the cohesive forces help to draw more water upwards.

At some point, the adhesive forces between the water and cellulose and the cohesive forces between the water molecules will be overcome by the gravitational forces on the weight of the water in the paper towel.  

When that happens, the water will not travel up the paper towel anymore. That is why it helps to shorten the length that colored water has to travel by making sure your paper towel isn’t too tall and making sure you fill your colored liquid to the top of the glass.

Rainbow Science Activity Extensions

Turn this demonstration into a true experiment by varying the water level (volume) you start with and seeing how long it takes the water to reach the empty glass.

Or start with the same volume of colored water and change the brand, type (single vs double ply, quilted vs not) or length of paper towel to see how long it takes for the water to “walk” to the empty glass.

You could even use the same volume of water, same length and brand of paper towel but vary the height of the filled glass , by raising them up on books, to see how that affects the speed of the water as it “walks” to the empty glass.

Have you had enough fun with the paper towels?  Try using other paper products to see how the type of paper effects the results.  Try toilet paper, printer paper, newspaper or a page from a glossy magazine.  What do you predict will happen?

Grab a Record Sheet

Help kids keep track of their results by grabbing our free record sheet! Then grab 30 more jaw-dropping (but easy prep!) science experiments kids will love from our shop!

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2- Ingredient Walking Rainbow Experiment That Works Like Magic

Categories Science Experiments

Kids will love making their very own walking rainbow experiment from just three colors as a fun addition to rainbow science experiments . It’s amazing how color mixing science experiments for kids can make something spectacular! The Walking rainbow science project is a classic science experiment that we’ve tried before. It was a lot of fun to watch the rainbow form right before our eyes!

This lesson teaches color theory, capillary action, the scientific method, and engineering. Kids will have a blast with this simple rainbow science experiment! It’s one of our favorite color science experiments for kids.

Table of Contents

What is the walking rainbow experiment.

This year, my kids realized they could make the entire rainbow with the walking water experiment starting with just the primary colors.

This rainbow walking water experiment packs a lot of learning into a tiny package! Kids will love watching the colors slowly transform over the course of 48 hours.

This project is also perfect for no prep STEM challenges , rainbow science experiments , St. Patrick’s Day STEAM activities , and more!

What do children learn from the walking water experiment?

The rainbow paper towel experiment   teaches a few basic scientific concepts.

When the paper towels are rolled up and placed between two jars, they exemplify capillary action, which is how liquid can move up something, rather than follow the usual pull of gravity and pull down.

To get a bit technical, intermolecular forces between the liquid and the paper towel creates surface tension that reacts with the adhesive force between the liquid and paper towel.

This causes the water to move up the paper towel and into the next jar.

What is the science behind climbing rainbow?

Capillary action is how plants pull water from the soil and up into their leaves to give them water and nutrients. This is the process that we will be mimicking in this rainbow walking water science experiment.

Then, once the paper towels pull color from the base red, blue, and yellow primary color jars, the resulting mixture creates the secondary colors of green, purple, and orange, completing the rainbow.

Water moves from jar to jar due to capillary action and surface tension. The surface tension of the water keeps the water from falling off the paper towel onto the table below as it creeps up the paper towel.

Capillary action is the force that is applied to the molecules in the water as they are absorbed by the towel. The pressure gently pushes the water all the way through the towel, and down into the jar next to it.

The water stabilizes and ends up at the same level in all the jars because of how things like to stabilize.

Once the water level is the same in all the jars and the paper towels are all wet, the water stops moving from jar to jar.

This is how the colors remain in primary in secondary colors, rather than continuously traveling from jar to jar, which would make all the jars turn brown.

STEM Classroom Decorations and Bulletin Boards

Here are some fun STEM themed classroom decorations and bulletin boards that will make any classroom more fun!

Cell Model Posters

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Walking Rainbow Experiment STEM Integration

Here are some additional STEM elements you can add to this activity to make it a complete STEM lesson plan!

Science: Color theory, capillary action. With paper towels (and plants work the same way), the molecules in the water are attracted to the molecules in the paper towels.

This causes the water to slowly move from the jar, up through the paper towel, and into the next jar. Eventually, the water level in all the jars will even out.

The way the walking rainbow experiment works is one of the most fascinating areas of science!

Technology: Food coloring is a marvel of modern science. We also used a timer to determine how long it took for the water to travel to one container to another.

Engineering: Kids have to set up the experiment properly or it won’t work. If you mix the wrong colors, you won’t end up with a rainbow, but a muddy-looking mess!

Math: Measuring, time estimation, comparison.

Walking rainbow experiment ingredients

Here is what you need to make this fun walking rainbow experiment.

The walking rainbow experiment teaches kids a lot about different parts of science, and this is what you need to make it happen!

• 6 mason jars
• Food coloring (in red, yellow, and blue)
• Paper towels

STEM Teaching Resources

These teaching resources will make your STEM classroom more fun and rewarding for your students!

STEM worksheets

STEM experiment kits

Space erasers

Planet erasers

Star erasers

Mini science notebooks

Science crazy straws

Science sticky notes

Science themed stickers

STEM journal

Life Cycle Puzzles

How long does the walking rainbow experiment take?

The walking rainbow water experiment does not take long.

In about 5 to 10 minutes, the color will move up the paper towel and start to fill the jar next to it.

Usually, the process is complete in just a couple of hours, although in some cases, it may take up to 8 hours for the colors to fully move from one jar to the other and combine to make the secondary colors.

Print the walking rainbow experiment directions!

Kids will love watching the new colors appear in this rainbow water experiment.

How to do the walking rainbow experiment

Kids will love making their very own walking rainbow from just three colors. This amazing walking rainbow experiment is the most fun walking water experiment ever! You'll have a blast with the rainbow walking water.

Instructions

Go over the color wheel before starting.

Show the kids how mixing colors will create different colors. However, don't tell them how to make the rainbow!

Let the kids discuss how they will make a rainbow from just three colors.

It will take them a little bit to determine the right combination.

If the kids are stumped, you can help them out by reminding them what colors are together in a rainbow.

The paper towels will start soaking up the water right away, but it will take about 48 hours before the process is finished.

Set a timer to find out exactly how long it takes to start mixing colors.

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Grow a Rainbow Science Experiment

Grow a rainbow with your kids! This simple experiment is so easy and fun! You just need markers, a paper towel and two cups of water! This experiment is quick to setup and just takes a few minutes to finish!

The great thing about this experiment is young kids can do almost all of it on their own! Our kids completed it several times! As always, make sure you supervise the kids and be sure to ask questions about the science experiment !

Supplies Needed:

• Two Clear Cups
• Marker (Washable Markers)
• Paper Towel

Video of Grow a Rainbow Experiment

Grow a rainbow instructions.

• Take a paper towel and fold it in half so it absorbs the water better. 
• You will then want your kids to measure the paper towel and cut it at around 7 inches. You don’t want it too long, as the colors might not connect in the middle if it is. Our cups were so short that the paper towel could have been a bit shorter and the colors would have met in the middle even better.

• You will want to use the markers to fill in about 6 or 7 rectangles on both sides of the paper towel. They should extend out about an inch and a half. Use the colors of the rainbow or close to it: Red, orange, yellow, green, blue, indigo, violet.

• After that, pour water into the 2 cups. You will want the cups to be about 1/2 to 3/4 full. This will depend on how tall your cups are.
• You will now gently put one end of the paper towel in one cup and the other end in the second cup – making a bridge between the two cups. Try to make sure the paper towel doesn’t fall too far into the water. If it does pull it out slightly or separate the cups a little more. 
• Now you just wait and watch the fun! The colors should connect in 5 – 10 minutes!

Additional Tips

• Do not place the paper towels too far into the water as the color will dissolve into the water and not travel as high.
• The more marker dye you use, the easier it will be for the colors to travel up and meet.
• Make sure you use a really washable marker. The more washable they are, the better the colors will travel up the napkins.
• Use an absorbent paper towel.

The kids had a blast with this science activity! They enjoyed trying different colors and amounts of marker. They would try to color in the sides of the paper towel with extra marker dye to make the colors connect even faster.

This activity definitely kept them busy and having fun. And it was easy to set up and quick to clean up too!

How does this simple science activity work?

In this rainbow experiment, capillary action is the reason the water moves up the paper towels.

Capillary action occurs because water is sticky since water molecules like to stay together (cohesion). And thanks to adhesion, water sticks to other substances and will move along their walls. This allows the water to move up the paper towels!

This same action is how water moves up into plants from the roots! It’s also why paper towels and pool towels are great at absorbing water!

Extend the Fun

Your kids can extend the fun by trying different colors, different paper towel lengths and different amounts of marker. Our kids tried to use a lot of marker to get the colors to grow faster!

They absolutely loved this science experiment! It was one of the easiest we have tried and they had a blast trying lots of variations!

Using markers, water and paper towels is a great way to show capillary action at work and amaze your kids!

Check out the Rainbow Walking Water and Color Changing Flower experiments below for more fun experiments showing capillary action.

More Fun Rainbow Science Experiments

The rainbow walking water is our most popular science activity! This one uses capillary action too!

You can use the colors of the rainbow with this color changing flower experiment ! This is another experiment using capillary action.

This  rainbow skittles experiment  is a fun rainbow activity for the kids!

Looking for more capillary experiments. This Celery Food Coloring Experiment is fantastic! – via Little Bins for Little Hands

Hello! I'm Laurence

Walking water experiment: make a rainbow using capillary action.

This walking water experiment is so easy and your kids will love it. It is a great way to teach them how capillary actions works.

What is Capillary Action?

Capillary action (sometimes called capillary motion or capillary effect) is the ability of a liquid to flow in narrow spaces without the assistance of, or even in opposition to, external forces like gravity.

This spontaneous rising of a liquid is the outcome of two opposing forces- cohesion (the attractive forces between similar molecules) and adhesion (the attractive forces between dissimilar molecules).

Why is capillary action important? Without capillary action, our blood wouldn’t circulate through our body. Trees wouldn’t be able to draw water into their roots and distribute it to their leaves. Simply put, life wouldn’t exist without capillary action.

What you will need for the walking water experiment:

• 5 clear glasses
• Blue, yellow and red food coloring
• Paper towels (4)

Fill 3 of the glasses with water. Add a few drop of blue food coloring into one jar, yellow in another and red in the third.

Arrange the glasses in a row so that they are in this order: blue glass of water, empty glass, yellow glass of water, empty glass, red glass of water.

Take a piece of paper towel and fold it so that it is in a skinny strip. Then connect the glasses with the folded paper towels.

The water will start moving quite quickly. The paper towel will draw the water upwards against gravity and then into the empty glass beside it.

Here is where you can teach a secondary lesson. Ask your kids what color they will  get if they mix blue and yellow. How about yellow and red?

As the colored water mixes into the empty glass, a new color will appear. When the water is finished “walking” there will be a beautiful rainbow!

This experiment moved a lot faster that I expected (about 20-30 minutes). Once the water levels are perfectly equal in each glass, the experiment is complete.

If it helps to watch a demonstration of this walking water experiment, check out this great example by Science Buddies:

This walking water experiment is such a fun and simple way to teach kids about capillary action. My kids loved making a rainbow of colors and even learned about mixing primary colors. Give it a try and let me know how it goes!

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One comment.

So interesting! Love the double lesson (capillary + color mixtures), and the simple beauty of this experiment!

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Walking Rainbow // Capillary Action

Introduction: Walking Rainbow // Capillary Action

Step 1: Materials

Step 2: Filling Glasses

Step 3: Food Coloring

Step 4: Paper Towels

Step 5: Wait!

Step 6: Compare With Real Life

Step 7: Watch It Live!

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Rainbow walking water science

Key points i covered in this post, what is the phenomenon behind rainbow walking water science, materials needed for the rainbow walking water experiment, setting up the experiment, understanding capillary action, the role of color mixing in the experiment, scientific observations and analysis, applications in the real world, is there a guide for conducting the rainbow walking water science experiment.

• Ensure you have all necessary materials: clear glasses, paper towels, water, and primary-colored food coloring.
• Arrange the cups in a circle or line, filling alternate ones with water and adding food coloring accordingly.
• Prepare the paper towels by folding them into sturdy strips and placing them to bridge between full and empty cups.
• Observe the water as it begins to “walk” up and across the paper towels and into the adjacent empty cups.
• Watch for the primary colors to mix, forming secondary colors and creating a full spectrum.
• Document any notable changes, such as speed and the total amount of water each paper towel can hold.
• Discuss the analogy to natural processes like plant water transport and other real-world applications of capillary action.

Can Walking Water Science Project be Conducted with Other Liquids Besides Water?

What colors are best for creating a rainbow effect in the experiment, how long does it take for the rainbow walking water to complete, is the rainbow walking water science project safe for young children, what scientific principles can be learned from this experiment, final thoughts, share article:, ocean slime recipe, summer science experiments.

Rainbow Celery: A Simple & Colorful Science Experiment

Hey, wonderful readers!

It’s Calie, back with another mesmerizing science experiment that’s sure to dazzle and educate both you and your kids. Today, we’re diving into the world of capillary action with an experiment that’s as visually stunning as it is educational – the Rainbow Celery experiment.

I remember finding this experiment online during a time when we were all searching for engaging activities to do at home. The idea of turning ordinary celery into a vibrant display of colors instantly caught my attention, and I knew it was something we had to try.

What You’ll Need

• Celery (with leaves for the best visual effect)
• Glasses of water
• Food coloring in various colors

Steps to a Rainbow of Learning

• Set the Stage : Find a sunny spot in your home where your celery can bask in the sunlight. This will not only help in the experiment but also make for a lovely display.
• Add Food Coloring : Fill each glass halfway with water and then add a few drops of different food coloring to each glass. For an extra touch of magic, arrange the colors in the order of a rainbow.
• Prepare Your Celery : Cut the stalks of celery so that they fit comfortably in the glasses without tipping them over. Place each stalk into a glass of colored water.
• Watch and Wait : This is where patience comes in. Let the celery stalks sit in the colored water overnight. It’s a slow process, but by the next day, you’ll start to see the leaves of the celery taking on the colors of the water they’ve been sitting in.

The Science Behind the Magic

This experiment is a fantastic way to introduce the concept of capillary action to children. Capillary action is the ability of a liquid to flow in narrow spaces without the assistance of, and often in opposition to, external forces like gravity.

The celery stalks, much like plants in nature, contain tiny “vessels” that transport water and nutrients from the roots to the leaves. In our experiment, these vessels also carry the colored water, allowing us to see how water moves through a plant.

By observing the celery change colors, children can get a visual understanding of how plants drink water and how capillary action is essential for their survival. It’s a simple yet profound demonstration that highlights the beauty and complexity of the natural world.

For those who want to see the process in action or need a little guidance, check out this helpful video: Rainbow Celery Experiment Video . It’s a great resource that can add an extra layer of engagement to this already fascinating experiment.

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100 Easy Science Experiments Your Kids Can Do At Home

Doing the Rainbow Celery experiment at home is not just about learning scientific concepts; it’s about creating memorable moments of discovery and wonder with your family. It’s a vivid reminder that science is all around us, waiting to be explored in the most unexpected places.

So grab some celery, a few glasses of water, and let’s bring the rainbow into our homes!

Happy experimenting, and here’s to filling our days with color and curiosity!

Warmest wishes,

Calie Herbst, Editor-in-Chief of Milwaukee With Kids, has spent over a decade combining her experiences as a parent of three to create a hub for Milwaukee’s family adventures.

Her decade-long teaching career in Milwaukee Public Schools and academic background, including a Master’s in Teaching from Marquette University and dual B.A.s in Sociology and Spanish from the University of Wisconsin – Madison, fuel her passion for inclusive and engaging family content.

Calie is also a recognized voice in local media, contributing to WISN Channel 12 News, WTMJ Wisconsin Morning News, Fox 6’s Real Milwaukee, and B93.3.

Discover more about Calie’s journey and editorial approach on her About Page  and Editorial Policy Page .

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Rainbow Walking Water

Discover the magic of color mixing and capillary action with this easy and exciting science project!

Ages: 3 - 8

Materials you'll need

• paper towel
• small cups (5)
• food coloring - red, yellow, blue

Step-by-step tutorial

Gather your materials.

Cut a few ½-inch wide strips from a piece of paper towel. Then, cut each strip in half lengthwise (aka hotdog style).

Place five cups about one inch apart from each other. Then, lay a small strip in between each of them.

Squeeze two drops of red food coloring into the cup on the far left, two drops of yellow food coloring into the cup in the middle, and two drops of blue food coloring into the cup on the far right. Leave the other two cups empty!

Pour water into each of the cups with food coloring, filling almost to the top.

Place one end of the paper towel into one color, and the other end into another. Watch what happens!

The color climbs up the paper towel bridges thanks to one nifty property of water — it likes to stick to things. (That’s why your finger gets wet when you dip it in water. The water molecules stick to you!) 

When water gets into a tiny space — like between paper fibers — it’ll stick to the sides of the fibers. Because it also sticks to other water molecules, more water gets pulled into the space after it. Gradually, by pulling in more water and sticking to the sides, the water will inch its way along and climb all the way up the paper. This process is called capillary action, and it’s also how water gets all the way from a tree’s roots up to the topmost leaves!

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Home Activity: Rainbow Capillary Action

Do you have a kid who is interested in science? Here’s an easy experiment on capillary action that even younger students can do. All you need are 7 cups, water, paper towels, and food dye in three colors: red, yellow, and blue. Arrange the cups close together in a line. Fold a paper towel into eighths. Cut off the ends so it fits into two of the cups, like a bridge. Repeat 5 more times so there are enough paper towels to connect all the cups, but do not leave them in the cups yet.

Starting with the first cup, fill every other one with water. Put red dye in the first water-filled cup, yellow in the second, blue in the third, and red in the fourth. Stir the water to make sure the dye is spread evenly throughout the water. Put the paper towels in the cups and wait.

Eventually, the dyed water will spread into the empty cups through the paper towels via capillary action. The colors will make the secondary colors in the empty cups, resulting in a rainbow throughout the row. This is because the water yearns for a state of equilibrium. Capillary action means that the liquid will spread evenly throughout the absorbent material, which is why the water spreads into the empty cups.

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Grow a Rainbow Experiment

Want to grow your own rainbow? Try this simple science experiment! You only need paper towel, water and washable markers. Kids will love to see their rainbow “grow” in this easy activity!

RELATED: Surprise Rainbow Activities 

You will love seeing the rainbow come together in this simple science experiment! You can even do different patterns and colors too.

Grow a Rainbow Experiment for Kids

Here is what you will need for this activity:  

• Paper Towel
• Washable Markers
• 2 Small Glasses

Watch the Full Video Tutorial Here

What is the science behind this experiment.

This science experiment is a great example of chromatography. Chromatography is a way of separating out a mixture of chemicals. If you ever got a paper with ink wet you would have seen the ink move across the page in streaks.

Capillary action makes the marker dye move up the paper towel.  The water moves upward through the paper towel, lifting the washable dye molecules with it. Because the washable markers are water based, they disperse in water.

Set up a few different scenarios and hypotheses. For example, if you were to try this experiment without any dye, you would still see the water rising upwards towards the center of the paper towel.

If you were try this experiment with permanent markers it would not work. This is because the markers are not water based (they are alcohol based) so the dye in the marker does not travel with the water. You can also show that permanent markers will disperse with rubbing alcohol but not with water.

• You need absorbent paper towel or napkin – we used the brand Bounty
• You must use washable markers – make sure to check it’s washable as not all Crayola brands are washable
• Do not place the end of the paper towel too deep into the water or the dye will dissolve into the water instead of traveling up the paper towel
• The shorter the paper towel – the better it works as there is less for the marker dye to have to travel across
• Add lots of marker to the ends.  You need lots of dye for it to travel upwards.

Growing Rainbow Experiment Instructions

1. Fold over a piece of paper towel (so you have 2 pieces on top of each other). Trim the length to be 7.5 inches (any longer and the rainbow may not connect fully).

TIP : The shorter your piece of paper towel, the better it will connect. Also make sure you are using an absorbent paper towel. We used Bounty.

2. Draw rectangles of the rainbow colors on each end.

You want to make sure to fill these colors in well so there is enough dye to travel across the paper towel.

TIP: Add lots of marker to the ends, you want a good amount of dye to travel up the paper towel.

3. Place 2 cups with water filled 3/4 full. You only want the bottom of the paper towel in so leave some space from the top of the cup.

Then place the paper towel into the cups, with one end in each cup.

TIP: Do not place the ends too deep in the water or the dye may dissolve into the water instead of moving up the paper towel.

4. The washable marker dye with slowly make it’s way up with the water to meet the other side in the center of the paper towel.

5. Leave the paper towel for 10-15 minutes and it will eventually connect the colors together.

We love how simple this science experiment is! Expand on the learning by testing with permanent markers or just water to see what happens in those scenarios.

More Rainbow Activities

Your kids will also love to try these surprise rainbow activities !

See this fun rainbow slime. Kids will love building a rainbow out of slime!

Make some rainbow playdough! This playdough recipe is super soft and lasts for months!

For another fun rainbow activity, this rainbow rice is our favorite sensory bin!

Related Ideas:

Cloud Dough

The BEST Playdough Recipe

50+ Christmas Crafts for Kids

Salt Dough Recipe

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8 comments on “grow a rainbow experiment”.

Me and my neighbors did the rainbow paper towel one and had lots of fun!

Hi can you pre color the paper towel? And do the experiment another day? We want to do it for a party and have it pre colored so all the kids have to do is dip it in water. 

Yes that should still work!

I love it. Thanks. I’m doing this as a virtual library lesson for a 25 minute class. If anyone is pressed for time like me it actually works faster with an unfolded cheap paper towel <5 minutes.

BEST experiment crafts I have seen in a long while. I hunt down ideas for my grandchildren when they come to visit and these are all on point and they will love them Thanks. I will use all of them (except the rice) . GREAT ideas.

Thanks so much Lee!

What is the purpose of folding the paper towel in half (step 1)?

We found that the colors moved across the paper better when there are 2 layers.

Walking Rainbow Experiment

You know when you attempt an experiment and it completely and utterly fails? Yeah, so that happened. But we were able to turn it into a fantastic study into scientific principles and the scientific method. In fact, it ended up being a much better lesson than if we hadn’t messed up. Yeah for failures! Wonder which experiment? It was the  very simple walking rainbow challenge . And I completely screwed it up ! Let me tell you more.

Rainbow Walking Water Science Experiment

What you will discover in this article!

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We are at the end of our school year. It’s been a HARD year! So I thought we would do a fun and easy science experiment that would bring a little color to our lessons and get the kids engaged and doing some fun activities. I chose the Walking Rainbow challenge.

Easy peasy lemon squeezy! All we had to do was make water walk. Seems easy enough!

Experiment Fail

Except I didn’t really do any research or read up on the activity. I was winging it. This was PRESCHOOL level stuff. So easy! I wanted a no prep STEM activity , but forgot that no prep doesn’t mean no research!

I grabbed 6 mason jars, Set them up in a little circle. Added equal amounts of water to each one. Then the kids added red to the first one, left one empty cup, yellow, skip, blue, skip. We talked about what colours were in the rainbow and learned the ROY G BIV (Red, Orange, Yellow, Green, Blue, Indigo, Violet) trick for remembering them.

Then we carefully folded up our paper towels, and placed them in our jars and waited for the magic to happen!

A few days later our colours had hit a major roadblock. Literally! The colored water defied gravity and climbed up the paper towels and then stopped. We waited a whole week and other than the colours getting more intense on the one side, nothing else happened.

A few days later our colours had hit a major roadblock. Literally, they climbed up and stopped. We waited a whole week and other than the colours getting more intense on the one side, nothing else happened.

Research Time

It was as if Gandalf himself was standing there declaring:

You shall not pass!!

Never one to give up, we sat down and studied what was happening in our rainbow walking water science experiment.

We learned that the experiment was about capillary action and the water travelling through the paper towels. We assumed, wrongly, that the primary colors would mix in the fold of the paper towel strips, and therefore mix in the various jars giving us our rainbow colors.

Instead the clear water and the coloured waters came up against each other and stopped. They didn’t mix in the paper towels.

Interesting… very interesting…

It was time to do some research and we learned that the middle jars where we didn’t add any food colouring, actually needed to be left empty. The mixing of the colors happens when the waters mix in the jars, not in the paper towels.

Rainbow Water Experiment Directions

We set up again. This time we did:

Cup #1 – a few drops of red food coloring and water Cup #2 – empty cup Cup #3 – a few drops of yellow food coloring and water Cup #4 – empty cup Cup #5 – a few drops of blue food coloring and water Cup #6 – finally another empty cup.

Then we folded our paper towel, making sure it wasn’t too long, but just long enough to touch the bottom of each jar.

We watched and waited. In a couple of days we had our rainbow colours! Red, orange, yellow, green, blue and purple!

Nature’s Balance

But we were curious, so we waited a whole week to see what would happen. We were rewarded with a wonderful demonstration of how the whole system brought itself into balance.

I talk to the kids a lot about the delicate balances in nature and this experiment provided the perfect visual representation.

After a week the water levels in all the jars balanced and became even. Each jar had exactly the same amount of water.

Science Explanation

The Walking Rainbow Experiment explores a few basic scientific concepts.

Capillary Action

At a basic level, when you fold paper towels and place them between two jars and one is filled with liquid, they demonstrate capillary action. A paper towel is made from cellulose fibers which have gaps. The water is able to travel through the gaps in the fibers. The water is able to move upward, against gravity, because of the attractive forces between the water and the fibers which act like capillary tubes.

Capillary action is how plants pull water from the soil, through the plant’s roots, and up into their leaves, we explored this concept in our Pollution Experiment .

For your older students, you can explore more technical details such as how the intermolecular forces between the liquid and the paper towel creates surface tension. This surface tension reacts with the adhesive force between the liquid and paper towel. This causes the water to move up the paper towel and eventually into the next jar. Where it eventually fills the second jar with water.

Dig into surface tension with our Magic Milk Experiment .

Science of Colour Mixing

The science of colour mixing is also part of this experiment. As the paper towels pull water molecules they also pull the food colouring from the original red, blue, and yellow primary color jars, it deposits that coloured water into the adjacent jar. This results in a mixing of the two colours from the adjacent jars, which gives us the secondary colours: orange, green, and purple.

Walking Water Rainbow

I set out planning on just bringing a little colour to our homeschool room, but this activity provided so much more. The learning and understanding for my kids went beyond a simple experiment and demonstrated something I want them to truly understand.

First: Nature wants to be in balance. It will work hard to find that balance. We need to do our best not to mess with that balance.

Second: Always do your research!

Also? It really was a beautiful experiment. And those colours? Stunning!

More Rainbow Experiments

5 Days of Smart STEM Ideas for Kids

Get started in STEM with easy, engaging activities.

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• Open access
• Published: 24 August 2024

The significance of electrical signals in maturing spermatozoa for phosphoinositide regulation through voltage-sensing phosphatase

• Takafumi Kawai   ORCID: orcid.org/0000-0001-6632-5551 1 ,
• Shin Morioka 2 ,
• Haruhiko Miyata   ORCID: orcid.org/0000-0003-4758-5803 3 ,
• Rizki Tsari Andriani 1 ,
• Sharmin Akter   ORCID: orcid.org/0000-0001-6901-3377 1   nAff9 ,
• Gabriel Toma 4   nAff10 ,
• Tatsuya Nakagawa 3 , 5 ,
• Yuki Oyama   ORCID: orcid.org/0000-0003-4579-5487 3 , 5 ,
• Rie Iida-Norita 3 ,
• Junko Sasaki 2 ,
• Masahiko Watanabe 6 ,
• Kenji Sakimura 7 ,
• Masahito Ikawa   ORCID: orcid.org/0000-0001-9859-6217 3 , 5 ,
• Takehiko Sasaki   ORCID: orcid.org/0000-0003-1837-3748 2 &
• Yasushi Okamura   ORCID: orcid.org/0000-0001-5386-7968 1 , 8  

Nature Communications volume  15 , Article number:  7289 ( 2024 ) Cite this article

Metrics details

• Cell signalling
• Phospholipids
• Reproductive biology

Voltage-sensing phosphatase (VSP) exhibits voltage-dependent phosphatase activity toward phosphoinositides. VSP generates a specialized phosphoinositide environment in mammalian sperm flagellum. However, the voltage-sensing mechanism of VSP in spermatozoa is not yet characterized. Here, we found that VSP is activated during sperm maturation, indicating that electric signals in immature spermatozoa are essential. Using a heterologous expression system, we show the voltage-sensing property of mouse VSP (mVSP). The voltage-sensing threshold of mVSP is approximately −30 mV, which is sensitive enough to activate mVSP in immature spermatozoa. We also report several knock-in mice in which we manipulate the voltage-sensitivity or electrochemical coupling of mVSP. Notably, the V312R mutant, with a minor voltage-sensitivity change, exhibits abnormal sperm motility after, but not before, capacitation. Additionally, the V312R mutant shows a significant change in the acyl-chain profile of phosphoinositide. Our findings suggest that electrical signals during sperm maturation are crucial for establishing the optimal phosphoinositide environment in spermatozoa.

Introduction

The membrane potentials of plasma membranes act as critical “electrical signals”, enabling neurons and other cells to communicate and transmit information. Therefore, the physiological implications of these signals have predominantly been explored in neurons or muscles, with a primary focus on voltage-gated ion channels that modulate ion permeability based on membrane potential 1 . In contrast, our previous work identified a distinctive voltage-sensing phosphatase (VSP) which shows the voltage-dependent phosphatase activity toward phosphoinositides (PIPs). This property is due to the unique architecture of VSP which has a voltage-sensor domain (VSD), phosphatase domain (PD) and VSD-PD linker 2 , 3 . VSD, which is also conserved in the conventional voltage-gated ion channels, consists of four transmembrane helical segments (S1-S4) and the S4 segment contains positive charge residues for sensing the membrane potentials. PD shows similarity to PTEN, which shows phosphatase activities toward PIPs. The VSD-PD linker connects PD with the VSD and plays crucial role in coupling VSD and PD activities. Thus, this unique configuration endows VSP with the capability to voltage-dependently dephosphorylate PIPs, thereby transforming “electrical signals” into “chemical signals” 3 , 4 , 5 , 6 , 7 .

Recently, our study revealed the functional expression of VSP in mouse spermatozoa, elucidating its role in regulating sperm motility during capacitation through analysis of VSP-deficient animal models 8 . VSP induced notable alterations in the PIPs profile of mature spermatozoa, creating a distinctive longitudinal PI(4,5)P 2 (Phosphatidylinositol 4,5-bisphosphate) gradient along the flagella. This specialized PI(4,5)P 2 environment emerged as crucial for modulating the activity of SLO3, a sperm-specific K + channel pivotal for fertilization 8 , 9 , 10 . However, several fundamental questions persist regarding the functional role of VSP in spermatozoa: Does VSP really sense the membrane potential of spermatozoa? If this is the case, what is the mechanism underlying it?

The voltage-dependent enzymatic activity of VSP was initially characterized in Ciona intestinalis (Ci-VSP) using Xenopus oocyte heterologous expression systems 2 . This activity was subsequently identified in vertebrates, including zebrafish (Dr-VSP), African clawed frogs (Xl-VSP), and chickens (Gg-VSP) in vitro 11 , 12 , 13 , 14 . However, attempts to detect voltage-dependent phosphatase activities using mammalian VSPs in heterologous expression systems have been unsuccessful, despite their structural homologies with non-mammalian VSPs 3 , 15 . Consequently, the existence of voltage-sensing capabilities in mammalian VSPs remains uncertain, contributing to a dearth of detailed information regarding the electrophysiological properties of these proteins.

Here, we started our study by investigating the PIPs profile at different stages of sperm maturation. Our findings reveal a progressive impact of VSP on the PIPs profile during sperm maturation, suggestive of continuous VSP activation throughout this long-term process. Additionally, two-electrode voltage clamp (TEVC) analysis of mVSP activity, with modifications to its intracellular linker and N-terminal sequence in a heterologous expression system, indicated that mVSP is activated at approximately −30mV, a membrane potential lower than the resting membrane potential of immature spermatozoa.

Finally, we generated three distinct knock-in mice expressing VSP mutations: (1) loss of voltage-sensing capability (D225R), (2) loss of electrochemical coupling between voltage-sensor and enzyme (K347Q), and (3) slightly altered voltage-dependency (V312R), based on the aforementioned TEVC experiments. While functionally null D225R and K347Q mutant mice abolished VSP expression itself, V312R mutant mice exhibited normal protein expression with significant changes in spermatozoa function. This study provides pioneering insights into the significance of voltage-sensing capabilities of mVSPs in native spermatozoa.

VSP demonstrates phosphatase activity during sperm maturation

Spermatozoa acquire fertility through a maturation process as they traverse from proximal to distal segments of epididymis, i.e., caput, corpus and cauda epididymis (Fig.  1a ). This process is pivotal for optimal sperm function. Notably, the lipid composition of spermatozoa undergoes changes during this process 16 . While the membrane phospholipids contain acyl groups, with two fatty acids located at the sn-1 and sn-2 positions (Supplementary Fig.  1 ), it remains unknown how the profile of acyl chains of PIPs transitions during the maturation process. To elucidate it, we employed liquid chromatography-tandem mass spectrometry (LC-MS/MS) to assess the profiles of PIPs as well as phosphatidylserine (PS) in both caput and cauda epididymal spermatozoa. This method allows for the measurement of both the carbon number and the number of double bonds of the combined acyl groups at the sn-1 and sn-2 positions (Supplementary Fig.  1 ), while it does not separate the isomers of PIP (PI(3)P, PI(4)P, PI(5)P) and PIP 2 (PI(3,4)P 2 , PI(3,5)P 2 , PI(4,5)P 2 ) (Supplementary Fig.  1 ). In line with previous findings 16 , we observed a marked difference in PS composition between caput and cauda spermatozoa (Supplementary Fig.  2a ). While PS contained only a limited set of acyl chain variants (Supplementary Fig.  2a ), PIP and PIP 2 in both caput and cauda spermatozoa exhibited diverse acyl chain variants, particularly enriched in Long-Chain Polyunsaturated Fatty Acids (LC-PUFA) (e.g., 40:5 and 40:6) (Supplementary Fig.  2b, c ). These LC-PUFA variants appear to include docosahexaenoic acid (DHA, 22:6) and docosapentaenoic acid (DPA, 22:5), which are highly prevalent in spermatozoa 17 , 18 . We confirmed the VSP protein expression at caput spermatozoa which is comparable to cauda spermatozoa (Supplementary Fig.  3 ). In cauda epididymis, the total PIP/PIP 2 ratio was significantly higher in mature sperm (cauda) than in immature sperm (caput) in VSP +/- samples (Fig.  1b ). This trend was absent in VSP-deficient samples, suggesting that VSP exerts its phosphatase activity during the maturation process.

a Spermatozoa, which differentiate in the testis, undergo maturation as they are transported through the caput epididymis to the cauda epididymis. By the time they reach the cauda epididymis, they become fully mature and are capable of fertilization. The matured spermatozoa are stored in the epididymal cauda. b PIP/PIP 2 ratio in spermatozoa from caput and cauda epididymis of each genotype mice. The significant difference of PIP/PIP 2 ratio between Vsp +/- and Vsp -/- was already observed in caput epididymal spermatozoa, but the difference is more pronounced in cauda epididymal spermatozoa (Tukey’s multiple comparison test. p -value is adjusted for multiple comparison. ** p  < 0.01, **** p  < 0.0001, n  = 3 independent mice for each group). The exact p -value is shown in the Data Source file. Data are represented as mean ± s.e.m. c – f PRMC-MS analysis was performed at different maturation stages of spermatozoa. The calculated PI(4)P/PI(4,5)P 2 ratios are shown. (unpaired two-sided t-test, ** p  < 0.01, *** p  < 0.01, **** p  < 0.0001). The data from cauda epididymis ( f ) was already reported in the previous study 8 . The experiment group in caput epididymis ( e ) corresponds to the experiment group of “low K + ” shown in Supplementary Fig.  4 . For ( c – f ), n  = 5,5,5 and 6 biologically independent mice in each genotype are used, respectively. Data are represented as mean ± s.e.m. The exact p -value is shown in the Data Source file.

Subsequently, we investigated whether short-term alterations in membrane potential influence the PIPs profile using sperm from the caput epididymis. We manipulated the membrane potential of caput epididymal spermatozoa by incubating them in high or low extracellular K + solutions with 10 μM valinomycin for 30 minutes and employed Phosphoinositide Regioisomer Measurement by Chiral column chromatography and Mass Spectrometry (PRMC-MS), which can accurately separate PIPs regioisomers 8 , 19 . No significant difference in PI(4)P/PI(4,5)P 2 ratio was observed between the treatments (Supplementary Fig.  4 ), suggesting that a short-time electric signal is not sufficient and prolonged VSP activation is required for exhibiting the effect of phosphatase activity.

Then, we conducted PRMC-MS measurements using spermatids and testicular spermatozoa, which are at the earlier maturation stages (Fig.  1c, d and Supplementary Fig.  5 and 6 , with cauda epididymis data included for comparison, as previously reported 8 , in Fig.  1f ). As shown in total PI(4)P/PI(4,5)P 2 ratio results (Fig.  1c–f ), the differences in the ratios between VSP +/- and VSP -/- gradually increase over this long-term maturation period. Furthermore, while spermatids exhibited no difference in total PI(4)P/PI(4,5)P 2 ratio between the genotypes, a significant difference was noted in LC-PUFA-containing variants (40:5 and 40:6) (Supplementary Fig.  6a , right ). Similarly, both in testicular spermatozoa and caput epididymal spermatozoa, a marked difference was evident for LC-PUFA-containing variants (e.g., 38:5, 38:6, 40:5, and 40:6) (Supplementary Fig.  6b, c , right ), with these distinctions diminishing in cauda epididymis (Supplementary Fig.  6d , right ). These results appear to suggest that mVSP may exhibit a preference for LC-PUFA during maturation, although the difference appears to be less obvious at the maturation stage, when the phosphatase activity is saturated.

mVSP activation occurs within the physiological voltage range of immature spermatozoa

Given that VSP demonstrates phosphatase activity during maturation, it is conceivable that the membrane potential of immature spermatozoa plays a crucial role in driving mVSP activity throughout its maturation process. We assessed the membrane potential of immature WT spermatozoa using the perforated patch clamp technique, preserving the intracellular environment. We observed that the averaged membrane potential of spermatozoa was −9.79 ± 1.23 mV ( n  = 15), consistently exceeding −30 mV in all recordings (Fig.  2a ). Subsequently, we sought to determine if mVSP could be activated within this range of sperm membrane potential. Considering its structural resemblance to other VSPs (Fig.  2b ), mVSP is anticipated to possess normal voltage sensitivity. However, as mentioned in the introduction, there are no successful reports demonstrating the voltage-dependent enzymatic activity of mammalian VSPs in vitro. This problem likely stemmed from the accumulation of mammalian VSPs in the Golgi apparatus of expression system cells, preventing their transportation to the plasma membrane 20 . Previous studies suggest that the N-terminal domains and intracellular loops of transmembrane proteins play crucial roles in trafficking to the plasma membrane 21 , 22 , 23 , 24 , 25 . Furthermore, while mVSP and Ci-VSP have an overall similar structure and conserved sequences, their sequences differ in the N-terminus and intracellular loops (Supplementary Fig.  7 ; full length: identity = 41.5%, similarity = 61.4%; N-terminus: identity = 18.9%, similarity = 40.0%). We addressed this by modifying mVSP, replacing the N-terminus and the intracellular S2-S3 loop with those of Ci-VSP, while preserving functionally critical regions: VSD, VSD-PD linker, and PD (Fig.  2c, d ). This modification significantly enhanced the surface expression of mVSP in Xenopus oocytes (Fig.  2e, f ). Here we refer to this molecule as mVSP*.

a Recording of the resting membrane potential from immature spermatozoa. n = 15 cells examined over 7 independent mice. Data are represented as mean ± s.e.m. b The structural comparison of VSD between mVSP (light blue) and Ci-VSP (orange). The VSD structure of mVSP was predicted with ColabFold, an opensource software for protein structure prediction 54 . The VSD of Ci-VSP was solved in the previous study 26 . Some important residues are shown with the numbering of mVSP amino acids residues. c Schematic diagram of the modified mVSP (mVSP*). The voltage-sensing domain, linker region for electrochemical coupling, and phosphatase domain remained intact. The sequence alignment of the VSD between mVSP and Ci-VSP is also presented with amino acids from the experiment shaded in blue and orange, respectively. d The structure of the mVSP* was predicted with AlphaFold2. mVSP and Ci-VSP regions are shown in cyan and orange, respectively. The N terminus region is omitted. e , f Immunoblots against the surface proteins and total protein were detected by HA-antibody. The signals ( e ) and statistics ( f ) are shown. (* p  < 0.05, unpaired two-sided t-test). N  = 5 independent samples in ( f ). Data are represented as mean ± s.e.m. The exact p -value is shown in the Data Source file. g Voltage-dependent regulation of KCNQ2/3 activities by mVSP*. KCNQ2/3 was co-expressed with either the original mVSP, mVSP* WT or mVSP* C458S, an enzyme dead mutant. A +50 mV depolarization pulse was applied to activate both the VSPs and KCNQ2/3. The holding potential is −60mV. In mVSP* WT, the KCNQ2/3 current gradually decreased. In contrast, the current decrease was not observed in KCNQ2/3 co-expressed with either the mVSP or mVSP* C458S. h Statistical analysis for percent reduction of KCNQ2/3 currents. There was significant difference between WT and C458S (Tukey’s multiple comparison test. p -value is adjusted for multiple comparison. *** p  < 0.001, **** p  < 0.0001). N  = 7, 8 and 8 independent experiments for mVSP, mVSP* and mVSP* C458S, respectively. Data are represented as mean ± s.e.m. The exact p -value is shown in the Data Source file. Voltage-dependency of mVSP* was analyzed using GIRK current. i Representative traces and the time course of GIRK current amplitudes with repetitive mVSP* activation. 1 s depolarization pulses (to activate mVSP*) were applied 21 times in the intervals of test pulses (−120mV, 100 ms). The red trace indicates the 21st trace. j Time course of the percent current reduction across repeated pulses. N  = 11 independent experiments. Data are represented as mean ± s.e.m. k The voltage-dependency of mVSP* was estimated from the percent current reduction at the 21st pulse in ( j ). N  = 11 and 3 independent experiments for mVSP* and mVSP* C458S, respectively. Data are represented as mean ± s.e.m.

We then evaluated the voltage-dependent phosphatase activity of mVSP* using KCNQ2/3 channels as a readout for PI(4,5)P 2 levels (Fig.  2g, h ). A notable decrease in KCNQ2/3 current occurred with prolonged depolarization-induced mVSP* activation, a phenomenon absents in non-modified original mVSP or enzyme-inactive mVSP* (mVSP* C458S), confirming its reliance on phosphatase activity. The voltage-dependency of mVSP* was further analyzed using GIRK (G protein-gated inwardly rectifying potassium channels) channels, applying repetitive 1-second voltage pulses with varying potentials (Fig.  2i ). The threshold for mVSP* activation was approximately −30 mV, and the activity gradually increased up to +50 mV, which was the upper limit of the repetitive 1-second long-pulse protocol, due to contamination from endogenous currents. Notably, the change in GIRK current amplitude was not observed in C458S. These results unequivocally demonstrate that mVSP can be activated above −30mV, a range within the physiological membrane potential of immature spermatozoa.

Detailed analysis of the voltage dependency of mVSP and its mutants

Given the upper limitation of +50 mV for the analysis with GIRK channels, an alternative technique was employed to scrutinize the detailed voltage-dependent properties of mVSP. We utilized PLCδ1-PH-GFP, a well-established PI(4,5)P 2 probe (Fig.  3a ), holding the membrane potential at −80 mV and applying a 10 s pulse protocol with different voltages up to +125 mV to examine the fluorescence change at the plasma membrane. Consistent with GIRK measurements, mVSP* activity was observed to begin around −30mV, reaching its maximum around +100 mV (Fig.  3b ).

a Schematic illustration of VCF experiments. b Left , Representative traces of fluorescence changes in mVSP*. 10 s pulses with different voltages were applied to observe fluorescence changes. The traces at various voltages are shown with different colors: black, cyan, blue, green, and red; to represent −50 mV, −25 mV, 0 mV, +50 mV and +100 mV, respectively. Right , The voltage-dependent activity of mVSP* WT as examined by fluorescence change. N  = 10 independent experiments. Data are represented as mean ± s.e.m. c Schematic diagram showing the mutated residues targeted in the VCF experiment. Representative traces of fluorescence changes in mVSP* mutants. d Effect of mutations on VSD. D225R, R309Q and V312R were examined based on previous studies. N  = 5, 6 and 7 independent experiments for D225R, R309Q and V312R, respectively. Because V312R showed significant difference from WT at −25 mV, it is also shown in the separate bar graph (Unpaired two-sided t-test, **** p  < 0.0001). Data are represented as mean ± s.e.m. The exact p -value is shown in the Data Source file. e Effect of VSD-PD linker mutation. K347Q did not show any fluorescence change. N  = 5. f Effect of PD mutation. C458S did not show any fluorescence change. N  = 4 independent experiments. Data are represented as mean ± s.e.m. Surface protein expression of mVSP and its mutants ( g , K347Q and D225R; h , V312R and C458S; i , R309Q) in Xenopus oocytes. Images (i) and statistics (ii) are shown. Unpaired two-sided t-test or Dunnett’s multiple comparisons were performed for comparison with WT, but there was no significant difference. N  = 5 independent samples in ( e ) and ( f ), while n  = 6 independent samples in ( g ). Data are represented as mean ± s.e.m. The exact p -value with adjustment for multiple comparison is shown in the Data Source file.

Subsequently, point mutation experiments were conducted (Fig.  3c ), based on the reported mutations in other species of VSPs to elucidate the similarities and differences of mVSP with its counterparts. We initially examined the effect of mutation on VSD (Fig.  3c and d ). The mutation on D225 in S1, considered a countercharge for Arg in S4 and essential for voltage-sensing capability 26 , was examined. In alignment with previous studies 27 , 28 , mVSP* D225R exhibited voltage insensitivity within the physiological range of membrane potential. Next, two distinct point mutations in S4, essential for the voltage sensing of voltage-sensor proteins, were induced: R309Q, generally demonstrating a large leftward shifted voltage dependency in diverse VSPs 11 , 14 , 29 , 30 , 31 , 32 , 33 , and V312R, exhibiting a moderate leftward shift in VSD activity of Dr-VSP (T156R) 11 . Unexpectedly, R309Q did not impact the voltage sensitivity of mVSP*, contrary to previous reports on other VSPs. On the other hand, V312R moderately but significantly changed the voltage dependency of mVSP* leftward at a lower voltage (Fig.  3c, d ), which is partly consistent with the observation in Dr-VSP 11 . These PLCδ1-PH-GFP results were corroborated in GIRK experiments (Supplementary Fig.  8 ), although differentiating between WT and V312R was challenging due to the limited information above +50 mV in GIRK recording. Additionally, a K347Q mutation in the VSD-PD linker region, critical for functional coupling 34 , abolished voltage-dependent phosphatase activity (Fig.  3c–e ). Furthermore, C458S, an enzyme dead mutant, displayed no VSP activity (Fig.  3c, f ). We also confirmed that surface expression of mVSP* mutants remained unchanged (Fig.  3g–i ).

In summary, these findings suggest that the fundamental mechanisms of voltage sensing and electrochemical coupling in non-mammalian VSPs are largely conserved in mVSP, with subtle differences in the voltage-sensing machinery involving R309 of S4 when compared to other VSPs.

The mVSP V312R mutation affects sperm function

Our findings suggest that mVSP undergoes activation driven by the membrane potential of spermatozoa throughout the entire maturation process. To explore the voltage-sensing capability of endogenous mVSP in sperm flagellum, we generated three distinct knock-in mouse models featuring the V312R (moderately altered voltage sensitivity; Supplementary Fig.  9a ), D225R (voltage-insensitive; Supplementary Fig.  9b ), and K347Q (no VSD-PD coupling; Supplementary Fig.  9c ) mutations, based on observations from in vitro experiments (Fig.  3 ). Functionally inert VSP mutants (D225R and K347Q) did not exhibit any VSP expression in spermatozoa (Fig.  4b ), implying that a certain level of VSP activation is necessary for maintaining its expression in spermatozoa. Conversely, the quantity of endogenous VSP protein in spermatozoa of homogenous VSP V312R mice was comparable to that of wild-type VSP (Fig.  4b ).

a Schematic diagram shows the mutated residues targeted in the knock-in mouse experiment. b Western blotting results show the protein expression of mVSP and Basigin (positive control) in native spermatozoa of WT ( Vsp WT/WT ), Vsp KO, V312R mutants ( Vsp VR/VR ), D225R mutants ( Vsp DR/DR ) and K347Q mutants ( Vsp KQ/KQ ). In Vsp DR/DR and Vsp KQ/KQ , the mVSP signal disappeared as well as Vsp KO. Unpaired two-sided t-tests were performed between the WT and homozygous mutants. ** p  < 0.01. For V312R, n  = 4, 5 and 1 independent mice for Vsp WT/WT , V312R mutants ( Vsp VR/VR ), and Vsp KO, respectively. For D225R, n  = 3 independent mice for each genotype. For K347Q, n  = 3 independent mice for each genotype. Data are represented as mean ± s.e.m. The exact p -value is shown in the Data Source file. c , d  Analysis of sperm motility before ( c ) and after ( d ) capacitation in WT and Vsp VR/VR . Left , Trajectories of spermatozoa isolated from WT and Vsp VR/VR mice. Spermatozoa were incubated for only 10 min in TYH in ( c ), while 2 h in ( d ). Right , Statistical analysis was performed on the percentage of cells showing circular motion using unpaired two-sided t -test (* p  < 0.05). N  = 5 independent mice for all experiment groups. Data are represented as mean ± s.e.m. The exact p -value is shown in the Data Source file. e Illustration of the parameters in sperm motility analysis. f , g  Quantitation of sperm motility parameters of non-capacitated ( f ) and capacitated ( g ) spermatozoa. The individual parameters are described in ( e ). Unpaired two-sided t -test * p  < 0.05. N  = 5 independent mice for all experiment groups. Data are represented as mean ± s.e.m. The exact p -value is shown in the Data Source file.

In a previous study, we found that VSP deficiency leads to abnormal sperm motility only after capacitation 8 . Therefore, we assessed the impact of the V312R mutation on sperm motility both before and after capacitation. We did not observe any differences in motility pattern between the wild-type and V312R homo spermatozoa before capacitation (Fig.  4c ). On the other hand, a substantial proportion of VSP V312R spermatozoa showed circular motion after capacitation which is significantly higher than wild-type littermates (Fig.  4d ). We also analyzed velocity using different parameters in detail (Fig.  4e ). Similarly, Vsp VR/VR spermatozoa exhibited a slight but significant difference in VCL (curvilinear velocity) and ALH (amplitude of lateral head) compared to Vsp WT/WT only after capacitation (Fig.  4g ), but not before capacitation (Fig.  4f ).

We also analyzed the PIPs profile in both immature and mature spermatozoa from the caput and cauda epididymis of Vsp WT/WT and Vsp VR/VR mice. Once again, the PI(4)P/PI(4,5)P 2 ratio progressively increased during sperm maturation in both Vsp WT/WT and Vsp VR/VR (Fig.  5a, b ). The total PI(4)P/PI(4,5)P 2 ratio remained nearly identical between Vsp WT/WT and Vsp VR/VR in both the caput and cauda epididymis (Fig.  5b ). However, LC-PUFA containing groups, especially 40:6, exhibited a trend of increase in Vsp VR/VR for both the caput and cauda epididymis (Fig.  5c–g ). In summary, the VSP V312R mutation, characterized by a leftward shift in voltage sensitivity, appears to exert a moderate but significant effect on spermatozoa function, underscoring the importance of the membrane potential in VSP function.

a Schematic diagram showing the V312R mutation residue and different maturation stages of spermatozoa. b Total PI(4)P/PI(4,5)P 2 analysis of spermatozoa in caput and cauda epididymis from WT ( Vsp WT/WT ) and V312R mutants ( Vsp VR/VR ). There was no difference between the two genotypes, although the PI(4)P/PI(4,5)P 2 ratio significantly increased during maturation in both genotypes. (Tukey’s multiple comparison test. p -value is adjusted for multiple comparison. **** p  < 0.0001, n  = 10 for each group). N  = 10 independent mice for each genotype. Data are represented as mean ± s.e.m. The exact p -value is shown in the Data Source file. c , d PI(4)P/PI(4,5)P 2 ratio profiles are identified based on sn-1 and sn-2 acyl chains in Vsp WT/WT and Vsp VR/VR spermatozoa from caput ( c ) and cauda ( d ) epididymis. n.a. indicates that the calculation cannot be performed due to no PI(4,5)P 2 detection. N  = 10 independent mice for each genotype, but the outlier is removed as appropriate. See also Source Data file. Data are represented as mean ± s.e.m. e Two-sided multiple-t test with no adjustments for multiple comparisons is performed and the p -value is shown in the table. The increases or decreases in Vsp VR/VR with a p -value < 0.05 were highlighted in red and blue, respectively. f , g The comparison of for (40:6) acyl chain in caput and cauda epididymis. (two-sided t-test., * p  < 0.05 and ** p  < 0.01, respectively. N  = 10 independent mice for each genotype. Data are represented as mean ± s.e.m. The exact p -value is shown in the Data Source file.

In this study, we have elucidated that VSP undergoes activation throughout sperm maturation, suggesting that the membrane potential of immature spermatozoa plays a crucial role in shaping the appropriate PIPs environment within the sperm flagellum (Fig.  6 ). This research marks the successful observation of the voltage-dependent enzymatic activity of mammalian VSPs in vitro, revealing that mVSP can be activated at the resting membrane potential of immature spermatozoa. Lastly, we discovered that the VSP mutant (V312R) with moderately shifted voltage-range-of-activation showed altered pattern of sperm motility and altered PIPs profile, further highlighting the importance of electric signal in maturing spermatozoa for VSP activation.

a mVSP activation during sperm maturation. Proper PI(4,5)P 2 environment of sperm flagellum, which is important for sperm function, is gradually formed during this process. b Voltage-dependency of mVSP in spermatozoa. We hypothesize that both V312R and WT are activated with resting membrane potential of maturing spermatozoa. D225R and K347Q shows protein degradation, because the basal phosphatase activity is required for VSP expression in spermatozoa.

Spermatozoa undergo dynamic lipid remodeling during epididymal maturation 35 . Here, we report that the PIPs profile is altered during sperm maturation, with Long-Chain Polyunsaturated Fatty Acids (LC-PUFA) preferentially incorporated into PIPs as spermatozoa undergo maturation. Furthermore, we identify VSP as a key player in this process, indicating that VSP is partly responsible for the lipid remodeling in terms of PIPs composition.

Considering that the PIPs level is generally balanced between phosphoinositide phosphatase and kinase activities, it is crucial to evaluate the contribution of kinases to this process. A previous study investigated the expression of phosphatidylinositol 4-phosphate 5-kinase (PIP5K) in spermatogenesis 36 . They reported that the expression of PIP5K isozymes (PIP5K1A and PIP5K1B) is highest in elongated spermatids but significantly reduced in epididymal spermatozoa. Consistent with this finding, our study reveals a gradual decrease in the PI(4)P/PI(4,5)P 2 ratio from spermatids to immature spermatozoa in VSP-deficient spermatozoa. However, this ratio does not undergo substantial changes in spermatozoa from the caput to cauda epididymis (Fig.  1b ). Overall, it is likely that PIP5K activity is relatively low in epididymal spermatozoa, with VSP emerging as the major regulator of PI(4,5)P 2 levels during epididymal maturation.

Notably, VSP exhibits a preference for targeting LC-PUFA during specific maturation stages (Supplementary Fig.  6 ). While the detailed molecular mechanism remains elusive, LC-PUFA, a newly incorporated lipid during maturation, maybe a susceptible target for VSP. Alternatively, our prior research unveiled that VSP establishes a heterogeneous distribution of PI(4,5)P 2 in the sperm flagellum 8 , which raises the possibility that LC-PUFA may also display heterogeneous distribution in the sperm flagellum, facilitating easy access for VSP to these molecules.

In this study, we demonstrated the voltage-sensing phosphatase activity of mammalian VSP using a heterologous expression system. Previous studies only reported the functionality of the PD of mammalian VSP by generating chimeras in which the enzyme domain was replaced from Ci-VSP to mammalian VSPs 12 , 15 , 37 . However, VSP requires other several factors such as VSD motion and VSD-PD coupling, in addition to enzyme activity, for proper function 3 . Therefore, it had been crucial to confirm the activity with the entire mammalian VSP structure. The present modified mVSP (mVSP*), retaining the intact VSD, VSD-PD linker, and PD, exhibited normal voltage-sensing phosphatase activity, strongly supporting the role of mVSP as a voltage-sensing phosphatase.

In heterologous expression experiments, mVSP* displayed phosphatase activity above −30mV. This threshold is comparable to that reported for Ci-VSP 2 , 3 and even more negative than other VSPs such as those from zebrafish, chicken, and frogs 3 , 11 , 12 , 13 , 14 . Additionally, mVSP* showed conservation of the common activation machinery shared with other non-mammalian VSPs. For instance, the mutation of D225 residue in S1 which is supposed to form a salt-bridge with Arg in S4 3 , 26 , 38 , completely abolished voltage-sensitivity consistent with the previous findings in Ci-VSD 27 . Furthermore, V312R mutation showed a moderate change in the voltage dependency, which is partially also consistent with the previous studies in Dr-VSP 11 . Similarly, the mutation K347Q in the VSD-PD linker of mVSP*, akin to other non-mammalian VSP 34 , demonstrated a common coupling mechanism between VSD and PD. Unexpectedly, the R309Q mutation in mVSP*, corresponding to R217Q in Ci-VSP, did not significantly alter voltage-sensitivity, contrary to previous reports for other VSPs 11 , 14 , 29 , 30 , 31 , 32 , 33 . This difference appears to suggest that the extracellular space of S4 in mVSP may have a distinct surface charge environment from other VSPs.

It is important to note that we modified the N-terminal and S2-S3 loop of mVSP for efficient expression in Xenopus oocytes. While this manipulation may potentially affect voltage-sensitivity, our previous study demonstrated that similar modifications of the N-terminal did not change the voltage-dependency of other VSPs or ion channels in Xenopus oocytes 21 , 22 . Also considering the lack of evidence that the intracellular S2-S3 loop critically regulates voltage-sensing phosphatase activity, despite comprehensive biophysical analyses of VSP 3 , it is unlikely that modifications to the N-terminal or S2-S3 loop of mVSP confer voltage sensitivity.

Our study suggests that VSP undergoes activation during epididymal maturation, a crucial process for establishing proper PIPs environments in matured spermatozoa. Therefore, we aim to discuss the voltage-sensing mechanism of VSP during this maturation process. Our heterologous expression experiments indicate that VSP becomes activated above −30 mV, a membrane potential range observed in immature spermatozoa. It is important to consider the possibility that the less negative membrane potential of immature spermatozoa could be also due to an incomplete pipette seal during the recording. However, when performing voltage-clamp experiments on the same measurements, we observed clear voltage-dependent currents (Supplementary Fig.  11a ), indicating that the quality of our measurements is reliable. Furthermore, alkalinizing the intracellular pH evoked a large hyperpolarization response in the same recording, which is consistent with previous papers (Supplementary Fig.  11b ) 9 . Therefore, it appears that the influence of leakage on our measurements of membrane potential is limited.

Notably, the several studies report that ion composition of the mammalian epididymal lumen significantly differs from the experimental HEPES-based solutions 39 , 40 , 41 , 42 . For instance, the rat cauda epididymis is reported to contain approximately 55 mM K + and only 20 mM Na + , potentially leading to a more depolarized state of spermatozoa compared to experimental conditions. Therefore, it is possible that extent of VSP activity in experimental condition was underestimated and VSP may be more efficiently activated at the resting membrane potential of epididymal spermatozoa.

Unexpectedly, single mutations in D225 (to R, causing voltage insensitivity) or K347 (to Q, resulting in no electrochemical coupling) eliminated VSP expression in spermatozoa (Fig.  4b ), although these mutants showed normal surface expression in heterologous expression systems. This implies that the basal phosphatase activity is essential for maintaining VSP expression in spermatozoa, and it is also consistent with our previous finding that functionally inactive Venus-tagged VSP lacked protein expression itself in spermatozoa 8 . Building on this idea, the observation that voltage-insensitive mutants (D225R) do not exhibit VSP expression appears to suggest that voltage sensing is crucial for VSP activity in spermatozoa.

Besides, V312R, which has leftward shift of voltage range for activation, showed modifications in PIPs profiles as well as sperm motility in the present study. Notably, the V312R mutant showed a differential trend in the LC-PUFA-containing PI(4,5)P 2 variant, possibly related to the observation that VSP selectively targets LC-PUFA at certain maturation stages (Supplementary Fig.  6 ). In the motility analysis, V312R exhibited the phenotype only after capacitation. This result is important because the sperm motility was significantly changed only after capacitation in VSP-deficient spermatozoa in the previous study 8 .

Previously, we observed that PI(4,5)P 2 levels are elevated in VSP-deficient spermatozoa, which enhances SLO3 activity and influences Ca 2+ signaling during capacitation 8 . It is also possible that some other regulatory mechanism by PI(4,5)P 2 is important for the capacitation, because the V312R spermatozoa only showed the partial increase in PI(4,5)P 2 levels in some acyl groups. For example, our previous study demonstrated that PI(4,5)P 2 is heterogeneously distributed along the sperm flagella with some clusters 8 , potentially affecting the localization of membrane proteins within the flagellum. Therefore, the altered PI(4,5)P 2 distribution might affect the protein localization that is important for capacitation. Interestingly, we observed that the expression of VSP itself is absent in mutants with severely impaired VSP function (D225R and K347Q). This result implies that the quantity of PI(4,5)P 2 potentially affect the expression of transmembrane proteins, including VSPs. It would be interesting, if VSP changes the expression profile of transmembrane proteins that are important for capacitation.

In conclusion, our results provide evidence for the critical role of voltage-sensing property of mVSP in spermatozoa. Future investigations into the regulatory mechanisms of membrane potential in maturing spermatids or spermatozoa will further enhance our understanding of the intricate relationship between membrane potential and enzyme activity.

In most experiments involving VSP-deficient animals, we utilized the same lineage as in our prior publication 8 . This lineage comprises VSP knock-in mice, where the PD was truncated at the active catalytic center, and Venus was fused at the end. Unexpectedly, this lineage lacked VSP protein in mature spermatozoa. Additionally, another VSP-deficient animal was generated to assess VSP protein expression in both immature and mature spermatozoa (See also Supplementary Fig.  3 ). All animal procedures were approved by the Animal Care and Use Committees of Osaka University.

Generation of another VSP-deficient mice

The pX330 plasmids expressing humanized Cas9 and single guide RNAs targeting exon 4 were injected into pronuclei of zygotes of B6D2F1 x B6D2F1 mice 43 , 44 . The sequence of gRNA is 5’-AGGTGTCAGTGAGTGCTTCA-3’. Embryos were cultured in KSOM overnight and subsequently transferred into the oviducts of pseudopregnant Institute of Cancer Research (ICR) outbred female mice. Screening of mutant pups was performed by direct sequencing following polymerase chain reaction (PCR) using primers (5’-ACCTGAAGCCATAGCTTAAGC-3’ and 5’-TTCTCCCACACTGGCTGGCTCAAG–3’). A founder mouse with an 1 bp insertion was used to expand the colony. The animal was backcrossed with C57BL/6 mice for three times. We confirmed that VSP protein expression is abolished in the homozygous mutant animals (Supplementary Fig.  3 ).

Plasmid and RNA Synthesis

mVSP was subcloned from mouse testis cDNA into the pSD64TF vector for cRNA synthesis. Hemagglutinin (HA)-tag sequence was introduced at the C-terminal of mVSP for detection in Western blotting. As illustrated in Fig.  2 , we replaced the N-terminal of mVSP (M1-S211) with that of Ci-VSP (M1-H115) and the S2-S3 loop of mVSP (V265-D276) with that of Ci-VSP (I169-N180) to design mVSP*. The nucleotide sequence of mVSP* is also shown in Supplementary Fig.  12 (see also Supplementary Table  1 for the primers). The number of amino acid residues in mVSP* (e.g., V312) is assigned with reference to the native mVSP. The GIRK2d (Kir3.2d) plasmid was provided by Dr. Yoshihisa Kurachi (Osaka University, Japan) 45 . G-protein β1 and γ1 subunit plasmids were provided by Dr. Toshihide Nukada (retired). KCNQ2/3 plasmids were provided by Dr. David McKinnon (Stony Brook University) and Dr. Koichi Nakajo (Jichi Medical University, Japan). Mutagenesis was performed using Primestar Max (Takara, Japan; see also Supplementary Table  1 for the primers). cRNA was synthesized using the mMESSAGE mMACHINE transcription kit (Thermo Fisher Scientific) after linearization with restriction enzymes.

Recordings of membrane potentials from immature spermatozoa

We performed perforated patch clamp recordings from immature spermatozoa to measure membrane potentials 8 , 46 . Sperm were isolated from the corpus epididymis in an HS-based solution containing (in mM): 135 NaCl, 5 KCl, 2 CaCl 2 , 1 MgSO 4 , 20 HEPES, 5 glucose, 10 lactic acid and 1 sodium pyruvate (pH 7.4). After 10 min, the supernatant was centrifuged, washed twice, resuspended in the HS-based solution, and placed on untreated glass coverslips. After 10 min, the coverslips were transferred to the recording chamber which was perfused with the HS-based solution. Recording pipettes were made of borosilicate glass (BF-150-86-10; Sutter Instruments, CA, USA) using a puller (P-97; Sutter Instruments). The intracellular solution contained (mM): 120 KCl, 3 MgCl 2 , 40 HEPES, 0.3 EGTA (pH 7.0), and 0.05 mg/ml gramicidin. We applied the negative pressure from the pipette to the cytoplasmic droplet of the spermatozoa to form the tight giga-ohm seal. After the giga-ohm seal, we waited for the pore formation by antibiotics which could be monitored by the access resistance. After pore formation, the access resistance was 50–100 MΩ. Recordings were performed using an Axopatch 200B (Molecular Devices, CA, USA) and sampled at 5 kHz using Digidata 1550 A (Molecular Devices) and pCLAMP 10.5 software (Molecular Devices).

Two-electrode voltage clamp recordings in oocytes

Xenopus oocytes were harvested from animals anesthetized in water containing 0.2% ethyl 3-aminobenzoate methanesulfonate salt (Sigma-Aldrich, St. Louis, MO). The oocytes were defolliculated by treating with type I collagenase (1.0 mg/mL; Sigma-Aldrich) in ND96 solution containing (in mM): 96 NaCl, 2 KCl, 5 HEPES, 1.8 CaCl 2 , and 1 MgCl 2 (pH 7.5). The defolliculated oocytes were then injected with cRNA. Current recordings were conducted two days after cRNA injection by two-electrode voltage clamp (TEVC) using an amplifier (OC-725; Warner Instruments, Hamden, CT). Acquired data were digitized using an AD/DA converter Digidata 1440 A running under pClamp at room temperature (22–24 °C). Output signals were digitized at 10 kHz. The bath solution was ND96, the glass electrodes were filled with 3 M KCl, and the resistances ranged from 0.2–1.0 MΩ. The holding potential was −60 mV.

Commercially obtained antibodies include goat polyclonal anti-BASIGIN (sc-9757, Santa Cruz, Santa Cruz, CA, USA), mouse monoclonal anti-β-Tubulin IV (T7941, Sigma-Aldrich), anti-HA (MMS-101R, Covance, Berkeley, CA, USA), Alexa Fluor 488-conjugated chicken anti-rat IgG (A-21470, Invitrogen, Carlsbad, CA, USA), Alexa Fluor 594-conjugated goat anti-mouse IgG (A-11005, Invitrogen), HRP-linked anti-rabbit or mouse secondary antibodies (NA9340V or NA9310V; GE Healthcare, Pittsburgh, PA, USA), and HRP-linked anti-goat secondary antibody (sc-2354; Santa Cruz Biotechnology). The VSP antibody used was the same as in our previous study 8 . The rat monoclonal anti-IZUMO1 antibody was generated previously 47 .

Isolation of Different Stages of Spermatids and Spermatozoa for PIPs Measurement

Mature and immature spermatozoa were isolated from cauda and caput epididymis, respectively. After making an incision in the epididymis, the tissues were stirred in an HS-based solution. After counting the cells, the sperm were centrifuged at 500  g for 5 min at 4 °C. The pellets were washed with PBS and centrifuged again at 500  g for 5 min at 4 °C. The pellets were then frozen and used for analysis.

For isolation of testicular spermatozoa and spermatids, we modified a previously reported two-step enzymatic digestion method 48 . Seminiferous tubules were dissociated with collagenase type I (1.0 mg/mL; Sigma-Aldrich) for 25 min at 32 °C in incubation medium; Hanks’ balanced salt solution (HBSS) supplemented with 20 mM HEPES (pH 7.2), 1.2 mM MgSO 4 , 1.3 mM CaCl 2 , 6.6 mM sodium pyruvate, and 0.05% lactate. Tubules were collected, and after a filtration step with a 40-µm nylon mesh, tubules were retained in the filter. The tubules were collected again and incubated at 32 °C for 25 min in the same collagenase buffer. The resulting whole cell suspension was filtered through a 40-µm nylon mesh to remove cell clumps. For testicular spermatozoa, the suspension was centrifuged at 300  g for 5 min at 4 °C and supernatant was collected. The sediment was used for spermatids collection as described later. The supernatant fraction was centrifuged with 800  g for 5 min at 4 °C, and then the supernatant was removed. After adding 1 mL incubation medium, it was centrifuged again at 800  g for 5 min at 4 °C. The pellet contained high purity of testicular spermatozoa (Supplementary Fig.  5b ). For isolation of spermatids, we used the above-mentioned sediment for flow cytometry and cell sorting as previously reported 48 , 49 . The cells were stained with Hoechst 33258 (5 µg/million cells; Dojindo, Kumamoto, Japan) for more than 30 min at room temperature. BD FACSAria IIIu was used for flow cytometry and cell sorting. Hoechst was excited using a 375 nm laser, and the dye’s wide emission spectrum detected in two distinct channels: “Hoechst Blue” (450/20 nm band-pass filter) and “Hoechst Red” (670 nm long pass filter). Forward Scatter (FSC-A) and Side Scatter (SSC-A) were detected using a 488 nm laser. The fraction of spermatids was collected in incubation medium (Supplementary Fig.  5a ) and centrifuged at 8000  g for 5 min at 4 °C. The cells were obtained from the pellet.

PIPs sample preparation

Mass spectrometric analyses of PIPs was performed as previously reported 19 . Frozen sperm samples were thawed and suspended in 1.5 mL of methanol. To this suspension, 50 µL of a methanol/chloroform (9/1) solution containing 1 nmol of C8:0/C8:0 PI(4,5)P 2 (serving as an absorption inhibitor) and 10 pmol each of synthetic C17:0/C20:4 phosphoinositide as internal standards were added. This mixture was then combined with 750 µL of ultrapure water, 750 µL of 2 M HCl, and 200 µL of 1 M NaCl. Following a thorough vortex-mixing, 3 mL of chloroform was added, and the mixture was vortexed again for 2 min. The resulting solution was centrifuged at 1200 g for 4 min at room temperature. The lower organic phase, which contains the crude lipid extract, was carefully collected and transferred into a fresh glass tube. PIPs were preconcentrated using an anion exchanging resin. DEAE Sepharose Fast Flow (10% slurry) was sequentially rinsed: twice with an equal volume of ultrapure water, once with 1 M HCl, twice again with ultrapure water, once with 1 M NaOH, and then twice with ultrapure water. The resin was then resuspended in methanol to create a 50% slurry, and a 0.5 mL bed volume was packed into a Pasteur pipette plugged with glass wool. The crude lipid extract (2.9 mL) was mixed with 1.5 mL methanol and applied to the column. The column was washed with 3 mL of a chloroform/methanol (1/1) solution, followed by 2 mL of a chloroform/methanol/28% aqueous ammonia/glacial acetic acid (200/100/3/0.9) solution. Elution was performed using 1.5 mL of chloroform/methanol/12 M hydrochloric acid/ultrapure water (12/12/1/1). The eluate was then mixed with 850 µL of 120 mM NaCl and centrifuged at 1200 g for 4 min at room temperature. The lower phase, containing purified PIPs, was collected into a fresh glass tube. The purified PIPs were derivatized through methylation, following the method of Clark et al 50 . Briefly, 150 µL of 0.6 M trimethylsilyl diazomethane was added to the purified phosphoinositide fraction prepared as described above at room temperature. After 10 min, the reaction was stopped by adding 20 µL of glacial acetic acid. The samples were then mixed with 700 µL of a methanol/ultrapure water/chloroform (48/47/3) solution, followed by 1 minute of vortexing. After centrifugation at 1200 g for 4 min, the lower phase was dried under a stream of nitrogen and redissolved in 100 µL of acetonitrile.

PI4P/PI(4,5)P 2 measurements by PRMC-MS

Phosphoinositide regioisomer measurement by chiral column chromatography and mass stepctrometery (PRMC-MS) was conducted using a QTRAP6500 triple quadrupole mass spectrometer (ABSciex) paired with a Nexera X2 HPLC system (Shimadzu) and a PAL HTC-xt (CTC Analytics) autosampler. Spectra were recorded in positive ion mode as [M + NH 4 ] + ions, with an MS/MS scan duration of 0.5 sec. The ion spray voltage was set to 5.5 kV, cone voltage to 30 V, and source block temperature to 100 °C. The curtain gas was set to 20 psi, collision gas to 9 psi, ion source gas pressures 1/2 to 50 psi, declustering potential to 100 V, entrance potential to 10 V, and collision cell exit potential to 12 V. Collision energy values for gas phase fragmentation are detailed in Supplementary Table  2 . A 10 µL lipid sample was injected using the autosampler, and lipids were separated using a CHIRALPAK IC-3 column (2.1 mmφ x 250 mm, 3 µm, DAICEL) in a 22 °C room. The liquid column chromatography was conducted at a flow rate of 100 µL/min with the following gradient: 40% mobile phase A (methanol/5 mM ammonium acetate) and 60% mobile phase B (acetonitrile/5 mM ammonium acetate) held for 1 minute, linearly increased to 85% mobile phase A over 2 min and maintained at 85% mobile phase A for 11 min. Data acquisition and processing were performed using Analyst 1.6.3 (SCIEX), while MultiQuant (SCIEX) was used for manual data evaluation and peak integration. No background subtraction was carried out, and Gaussian smoothing width was set to 1.0 points. For quality control, peaks from samples where the cps of the surrogate internal standards (SIS; C37:4 PIPs) from the multiple reaction monitoring (MRM) scan were below 2 × 10 4 were excluded from quantification analysis. The sample peak area value was divided by the corresponding SIS peak area value (equivalent to 1 pmol) to achieve relative quantification. Supplementary Table  3 lists the MRM transitions (pairs of m/z values of precursor ions and fragment/diacylglycerol ions) used for the identification and quantification of each PI4P/PI(4,5)P 2 molecular species.

Reverse phase (RP) LC-MS/MS analysis

An Ultimate 3000 LC system (Thermo Fisher Scientific) was used for the RP LC-MS/MS analysis, connected in tandem to a TSQ Vantage triple stage quadrupole mass spectrometer (Thermo Fisher Scientific) operating in positive-ion mode. The derivatized phospholipids (the injection volume was 20 μL, and the flow rate was set at 220 μL/min) were separated on an InertSustainBio C18 column (GL Sciences) with the following solvent gradient: 0–1 minute hold at 70% A/30% B, 1–3 min linear gradient to 90% A/10% B, 3–7.5 min constant at 90% A/10% B, and 7.5–13 min at 30% A/70% B. Here, mobile phase A consisted of acetonitrile/ ultrapure water/70% ethylamine (800:200:1.3), and mobile phase B consisted of isopropanol/acetonitrile/70% ethylamine (800:200:1.3). Measurement of PIP and PIP 2 species was achieved through MRM using a pre-set list of mass to charge ratio values (Supplementary Table  4 ). Spray voltage was set to 3250 V, sheath gas pressure to 15 arbitrary units, capillary temperature to 300 °C. Supplementary Table  5 lists the voltage for fragmentation. Data acquisition and processing and peak integration were performed using Xcalibur software 2.0 (Thermo Fisher Scientific). The quantification was achieved by dividing the sample peak area value by the corresponding the internal standard peak area value.

Western blotting from spermatozoa

Sperm were isolated from caput and cauda epididymis. They were rotated at 4 °C for 1 hour in a lysis solution containing: 10 mM Tris-HCl (pH 7.5), 50 mM KCl, 1% Triton X-100, and cOmplete™ Protease Inhibitor Cocktail (Roche). After centrifugation (9000 g for 5 min at 4 °C), the supernatant was mixed with sample buffer and 2.5% 2-ME. After SDS-PAGE, the proteins were transferred to a PVDF membrane. After blocking with 0.5% skim milk or 2% BSA, the blots were incubated with the primary antibody: anti-VSP (1:500) or anti-BASIGIN (1:500) in Can get signal 1 (Toyobo, Osaka, Japan). The membranes were washed and incubated with HRP-linked anti-rabbit or goat antibody (1:1000) in Can get signal 2 (TOYOBO). The signals were detected with ECL Prime Western Blotting Detection Reagent (GE Healthcare). Images were acquired using a CS analyzer system (ver. 3) (ATTO, Tokyo, Japan). Antibody stripping was sometimes performed using 200 mM Glycine (pH 2.8) for 10 min at 60 °C for other antibody experiments. The signal intensity of individual bands was calculated by subtracting the background intensity.

Western blotting of membrane proteins from Xenopus oocytes

Two days prior to the experiment, Xenopus oocytes were injected with cRNA. They were then placed in ND96 solution containing EZ-Link Sulfo-NHS-SS-Biotin (0.5 mg/mL, Thermo Fisher Scientific) for 30 min at room temperature. Cells were washed with PBS three times and quenched with lysed in 300 µL PBS supplemented with Triton X-100 (1%; Sigma Aldrich) and Complete protease inhibitor cocktail tablets without EDTA (Roche, Basel, Switzerland). After centrifugation (15310  g for 10 min at 4 °C), 150 µL of the supernatant was retained as “total lysate,” and the rest was incubated overnight at 4 °C under gentle rotation with 30 µL streptavidin agarose beads (COSMO BIO, Tokyo, Japan) pre-washed with PBS. The beads were collected by centrifugation (15310  g for 10 min at 4 °C) and washed with PBS containing 1% Triton X-100 three times. Biotinylated proteins were eluted from the streptavidin agarose beads by incubating in SDS-PAGE sample buffer containing 2.5% 2-ME for 30 min at room temperature (“Cell surface”). Total lysates were also mixed with SDS-PAGE sample buffer with 2.5% 2-ME. After SDS-PAGE for the supernatant, the proteins were transferred to a PVDF membrane. After blocking with 0.5% skim milk the blots were incubated with the primary antibody: anti-HA (1:2000, Covance, Berkeley, CA) in Can get signal 1 (Toyobo, Osaka, Japan). The membranes were washed and incubated with HRP-linked anti-mouse antibody (1:2000, Cytiva, Massachusetts, USA) in Can get signal 2 (TOYOBO). The signals were detected with ECL Prime Western Blotting Detection Reagent (GE Healthcare). Images were acquired using a CS analyzer system (ver. 3) (ATTO, Tokyo, Japan).

Sperm motility analysis

Sperm velocity was analyzed as described previously 51 . Spermatozoa isolated from the cauda epididymis were suspended in TYH medium, a well-established capacitation-inducing medium 52 . TYH contains: 120 mM NaCl, 4.8 mM KCl, 1.2 mM KH 2 PO 4 , 5.6 mM glucose, 1.0 mM sodium pyruvate, 1.7 mM CaCl 2 , 1.2 mM MgSO 4 , 25 mM NaHCO 3 , 4.0 g/L ALBMAX I (Thermo Fisher Scientific), penicillin (50 units/mL)-streptomycin (50 μg/mL), and 0.6% Phenol-red. Average path velocity (VAP), curvilinear velocity (VCL), straight-line velocity (VSL), and Amplitude of Lateral Head Displacement (ALH) were measured using the CEROS II sperm analysis system (Hamilton Thorne Biosciences, MA, USA) at 10 min and 2 h after incubation. Sperm motility was videotaped with an Olympus BX-53 microscope equipped with a high-speed camera (HAS-L1, Ditect, Tokyo, Japan) at 200 frames per second. The trajectory was visualized with 450 frames using ImageJ software (NIH) and plug-in Color Footprint Rainbow developed by Y. Hiratsuka (JAIST, Ishikawa, Japan).

Voltage Clamp Fluorometry (VCF) recording with PHPLCδ1-GFP

The plasmid of the GFP-fused pleckstrin homology domain from the PLCδ1 subunit (PHPLCδ1-GFP) was used as previously reported 53 . The microscope BX50WI upright fluorescence microscope (Olympus, Japan) was used with a 20 × 0.75 N.A. objective lens and LED lamp (MCWHL8: Thorlabs, Inc., New Jersey, USA), fitted with an excitation filter of BP460-480HQ (Olympus) and an emission filter of BA495-540HQ (Olympus). The emitted light is detected by a PMT (H10722-20; Hamamatsu Photonics, Japan). TEVC recording was done using the amplifier, Oocyte Clamp OC-725C (Warner Instruments, USA). Data were digitized using Digidata 1440A (Molecular Devices, USA) run by the software pClamp 10.3 (Molecular Devices, USA) with a 10 kHz sampling rate on Windows PC. After digitization, data were digitally filtered at a cut-off frequency of 50 Hz on pClamp. The bath solution was ND96, and the glass electrodes were filled with 3 M K + -acetate and 10 mM KCl. The resistances ranged from 0.2-1.0 MΩ. The holding potential was −80 mV. A depolarizing pulse (−50 to 125 mV) was applied for 10 s.

Generation of VSP point-mutation knock-in mice

In this study, we generated D225R, V312R, and K347Q mice. C57BL/6 fertilized eggs were obtained by in vitro fertilization. Oligonucleotides (200 ng/μl) and crRNA/tracrRNA/Cas9 ribonucleoproteins (40 ng/μl crRNA plus tracrRNA, 100 ng/μl CAS9) were electroporated into the fertilized eggs using a super electroporator NEPA21 (NEPA GENE, Chiba, Japan) (poring pulse, voltage: 225 V, pulse width: 2 ms, pulse interval: 50 ms, and number of pulses: +4; transfer pulse, voltage: 20 V, pulse width: 50 ms, pulse interval: 50 ms, and number of pulses: ±5). The reference oligonucleotides contained the point mutations D225R, V312R, and K347Q, respectively. Furthermore, it contains silent mutations that prevent recutting of the target sequences and allow recognition by the restriction enzyme for convenient genotyping (Fig.  4 and Supplementary Fig.  6 ). The reference oligonucleotides are listed in Supplementary Table  6 . One out of 25, 8 out of 24, and 5 out of 15 pups contained the expected mutations in D225R, V312R, and K347Q, respectively. The animals were crossed with C57BL/6 mice to expand the colony. The genotype primers for each knock-in mouse are as follows: D225R, 5′–GGGGCTTGGTGCATACTTTA–3′ and 5′–AGCTGTGACAAAGCCACTG–3′; V312R, 5′–CATTGCCCTTTGTCTTCTAC–3′ and 5′–ATTGGGAAATCATAAAGCTG–3′; K347Q, 5′–CCTTGTGTCTCGGGGAAATA–3′ and 5′–GCTCACGTGACTCAGGGAAT–3′.

Immunocytochemistry for isolated spermatid and testicular spermatozoa

The isolated spermatids or spermatozoa in incubating medium were seeded on 1 mg/mL poly-L-lysine coated coverslips. After 1 h, the sample was fixed with 4% PFA/PBS. Cells were washed with 0.3% PBST. For the primary antibody, a rat monoclonal anti-IZUMO1 (1:500 dilution) and anti-β-Tubulin IV (1:500 dilution) was used with 10 % goat serum. The binding of the primary antibody was detected using Alexa Fluor 488-conjugated chicken anti-rat IgG (1:2000 dilution) and Alexa Fluor 594-conjugated goat anti-moue IgG (1:2000 dilution). Confocal images were acquired using a LSM770 confocal laser scanning system (Carl Zeiss, Germany).

Litter size analysis

Male mice of each genotype were crossed with 8 − 11 weeks old of wild-type female mice. The number of pups was defined as litter size.

In vitro fertilization

Mature cumulus intact oocytes were collected from wildtype females and placed in a drop of 100 μl TYH medium. Spermatozoa were collected from cauda epididymis of male mouse and incubated in TYH medium for 2 h to induce capacitation. Capacitated spermatozoa were added to the TYH drop containing oocytes at a final concentration of 2 × 10 ^5 sperm/ml. Embryos reaching the two-cell stage by the next day were counted as fertilized.

Data Analysis

Data analysis was performed with Excel 2016 (Microsoft, USA), Clampfit 10.5 (Molecular Device, USA), and Igor Pro 6.37 (WaveMetrics, USA) software. Statistical analysis was performed with Prism 6 (GraphPad Software, San Diego, CA). For two-group comparison, we conducted an unpaired t-test or Mann-Whitney test as appropriate. An outlier was detected with Grubbs’ Test (α = 0.05) in individual experiment groups and removed from the analysis (Fig.  5c, d ). For multiple comparisons, we conducted multiple t -test, Dunnett’s multiple comparisons or Tukey’s test as appropriate. Data are represented as mean ± s.e.m. *, **, ***, and **** indicate a significant difference: p  < 0.05, p  < 0.01, p  < 0.001, and p  < 0.0001, respectively.

Reporting summary

Further information on research design is available in the  Nature Portfolio Reporting Summary linked to this article.

Data availability

The data generated in this study are provided in the main text or the Supplemental materials.  Source data are provided with this paper.

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Acknowledgements

We express our gratitude to Ms. Hikari Ginama, Ms. Megumi Kobayashi, and Dr. Natsuki Mizutani from Osaka University, Dr. Hiroki Nakanishi from Akita University, and NPO Biotechnology Research and Development for their technical support. We also thank Ms. Risa Mori-Kreiner for critical reading of the manuscript. This study received support from the Center for Medical Research and Education at the Graduate School of Medicine, Osaka University, and JSPS KAKENHI Grant Number JP20KK0376 (AdAMS) to T.K. Funding was provided by Grants-in-Aid from JSPS KAKENHI Grant Numbers JP17K15558, JP20K07274, JP20KK0376, JP23K06334 and JST FOREST Program, Grant Number JPMJFR225Z. Additionally, financial support was received from The Ichiro Kanehara Foundation, the Hyogo Science and Technology Association, the Sumitomo Foundation, the Ono Medical Research Foundation, the Uehara Memorial Foundation, the Senri Life Science Foundation, the Takeda Science Foundation, and Mochida Memorial Foundation for Medical and Pharmaceutical Research, all of which contributed to T.K.ʼs research. T.S. was supported by AMED under Grant Number 24gm1710007, by TMDU under Multilayered Stress Diseases (JPMXP1323015483), and Medical Research Center Initiative for High Depth Omics. This work was also supported by MEXT KAKENHI Grant Numbers JP15H05901, JP20H05791) and JSPS KAKENHI Grant Numbers JP21229003, JP25253016, JP19H03401 to Y.Okamura.

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Sharmin Akter

Present address: Department of Physiology, Bangladesh Agricultural University, Mymensingh, Bangladesh

Gabriel Toma

Present address: Graduate School of Frontier Biosciences, Osaka University, Suita, Japan

Authors and Affiliations

Graduate School of Medicine, Osaka University, Suita, Japan

Takafumi Kawai, Rizki Tsari Andriani, Sharmin Akter & Yasushi Okamura

Department of Biochemical Pathophysiology/Lipid Biology, Medical Research Institute, Tokyo Medical and Dental University, Tokyo, Japan

Shin Morioka, Junko Sasaki & Takehiko Sasaki

Research Institute for Microbial Diseases, Osaka University, Suita, Japan

Haruhiko Miyata, Tatsuya Nakagawa, Yuki Oyama, Rie Iida-Norita & Masahito Ikawa

Center for Medical Research and Education, Osaka University, Suita, Japan

Graduate School of Pharmaceutical Sciences, Osaka University, Suita, Japan

Tatsuya Nakagawa, Yuki Oyama & Masahito Ikawa

Faculty of Medicine, Hokkaido University, Sapporo, Japan

Masahiko Watanabe

Brain Research Institute, Niigata University, Niigata, Japan

Kenji Sakimura

Graduate School of Frontier Bioscience, Osaka University, Suita, Japan

Yasushi Okamura

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Contributions

T.K. and Y.Okamura conceived the study; T.K. and Y.Okamura designed experiments; T.K., H.M, Y.Oyama, and R.I.N performed and M.I. supervised experiments for in vitro fertilization and sperm motility analysis; T.K, S.A. and G.T performed FACS for spermatids isolation; S.M. performed experiments for LC/MS-MS analysis and J.S and T.S supervised them; T.N and H.M generated knock-in mice and knock-out mice; K. S provided VSP-deficient mice; M.W provided the antibody for VSP; T.K and R.T.A performed VCF experiments; T.K performed experiments for immunocytochemistry, Western blotting, and electrophysiology. T.K. drafted the manuscript; All members critically read the manuscript.

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Kawai, T., Morioka, S., Miyata, H. et al. The significance of electrical signals in maturing spermatozoa for phosphoinositide regulation through voltage-sensing phosphatase. Nat Commun 15 , 7289 (2024). https://doi.org/10.1038/s41467-024-51755-2

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DOI : https://doi.org/10.1038/s41467-024-51755-2

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