chlorophyll experiment leaves

Embrace the gorgeous colour of fall this year by conducting a science experiment that shows the colours leaves turn in the fall and explore the science behind this gorgeous transformation that occurs every autumn. We love a simple science experiment and this Leaf Lab is perfect for fall as we explore the big question: Why do leaves change color? Kids from preschool to high school will enjoy this easy and fun Fall science experiment!

Fall Science Why Do Leaves Change Color in the Fall?

What you will discover in this article!

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I have a love-hate relationship with fall. I love the gorgeous colours and the crispness of the air. And don’t forget all the amazing freshly harvested fruits and vegetables.

But there is also the dread that means winter is coming, and when you live in a place with brutally cold temperatures and snow up to 8 months of the year, it makes the signs of it’s arrival, a little panic inducing.

It’s like a warning sign… GET OUTSIDE NOW!

That’s what makes this Fall Science Experiment so perfect. Not only do you get to learn something fascinating about nature, like exploring how leaves change color, but you also get to spend some time outside!

I highly recommend doing this experiment just as the leaves start changing, while some are still green, and others are at various stages of changing colours.

LEARNING EXTENSIONS EXPLORE MORE FALL STEM ACTIVITIES

Leaf Science Chromatography Experiment

When I first looked into this activity, I came across some sites claiming that this activity would reveal what colours the leaves turn in the fall. After doing a few tests we knew that claim was not accurate. Instead, what this activity does is separate out the active colours in the leaves. By active, we mean the colours need to be at least a little bit visible in the leaf. That means for the best results, try picking handfuls of leaves from the same tree but that are at different stages of changing colours. We have a Mountain Ash that gave us leaves in 4 different colour groupings.

Or you can pick different leaves, from different trees, but don’t expect colours to magically appear that are not visible already in the leaf with this simple science approach.

For a more advanced type of Leaf Chromatography experiment for your middle school and high school students, check out this Leaf Lab . We haven’t done this technique but it looks like it might provide a slightly different result from this more simple science, Leaf Chromatography experiment. And may reveal more of the hidden colour pigments in the leaves.

Our Experimental Method

With this experiment we started by doing the steps with leaves from different trees on our property. These are trees we know very well. Beloved trees we have watched move for the seasons for at least a decade. So we are very familiar with their fall colours.

That’s how we discovered that this experiment doesn’t show what colour the leaves will turn in the fall, and therefore doesn’t really answer our big question: Why do leaves change color? Our tree that we know turns a gorgeous red in the fall, did not show any red at all in the chromatography.

But interestingly, our purple leaf tree, that starts in the spring with green leaves, that turn purple in summer, then orange/yellow in fall, gave us the most dynamic chromatography. We even made a cool heart chromatography!

So our next step in the study we went to our Mountain Ash tree. A HUGE very loved, majestic part of our yard. It also possesses the most amazing array of colours as it enters fall. Right now it has green, yellow-green, purple-green and a gorgeous bright berry red. So many beautiful leaf colours in one massive tree!

Then we redid the experiment. If it was true that the experiment would reveal what colours would appear in the fall, then all the chromatography should have revealed the same colours since the leaves were all from the same tree, picked at the same time, just at different stages of turning to the fall colours.

It turns out that wasn’t the case at all. It was only the active colours that were revealed. Which was a great finding! Which turned this Leaf Lab into an amazing journey of discovery and the scientific method .

To do this fall leaf experiment you will need:

Leaves ! See tips above for the best leaf selection Jars with lids (or plastic wrap will work if you don’t have lids) Rubbing Alcohol Butter knife (or other item that can “mash leaves”) Large bowl or dish that fits the jars Hot water Coffee filters Plates Scissors

Time needed:  2 hours and 30 minutes

Reveal the secret colours of leaves with this science experiment.

Start by heading outside and collecting leaves. I used small bags to keep my leaves organized. In total we did 6 different types of leaves. You want to gather 4 to 6 leaves of each type. Try and get leaves that are at different stages of turning colour for the best results.

Set out one jar for each type of leaf you collected. Into each jar start ripping up one type of leaf. You want to tear the leaves up nice and small. Really tear it up!

Add just enough rubbing alcohol to cover the leaves, then using a knife or other utensil mash up the leaves into the alcohol even more. You should start to see the rubbing alcohol turning a bit green.

Set the jars in a large dish or bowls. Add hot water to the bowl or dish and cover the jars with lids or plastic wrap.

Now let them sit for approximately one hour. Give the jars a little swish every once in a while to help release the chlorophyll. Your alcohol should be turning a nice green. You can also refresh the hot water if it cools too much.

Cut a coffee filter into quarters and place on a plate. Using a teaspoon or dropper add a couple of drops of the liquid to the tip of the filter.

Wait approximately 30 min to one hour to see how the colours move up the filter paper and separate. Pretty cool!

What happens if you leave it in the solution even longer? Maybe overnight? The colours will become more bold and larger, making the colours easier to see.

The Science Of Leaf Color Changes In Fall

Let’s dig into the big question: Why do leaves change color?

Why are Leaves Green?

Leaves, like all green plants, contain Chlorophyll. Chlorophyll is the molecule that traps the energy of the sun and is called a photoreceptor. It is found in the chloroplasts of green plants, and is what makes green plants, green. It is also a key part of photosynthesis which is what allows plant to get energy from sunlight.

Leaf Coloring

Leaves are actually more colours than just green but Chlorophyll is so dominant most of the summer that its green colour covers up all the other natural colours in the leaves. We can separate the colours of a leaf by doing this leaf chromatography science experiment.

We have done chromatography experiments before, check out our gorgeous Circuit Chromatography Flowers that light up!

In this science experiment we used rubbing alcohol and two types of energy to separate the colours. We used mechanical energy by ripping up the leaves and mashing them, plus heat energy from the hot water.

Look closely at the rubbing alcohol, do you see any other colours in there?

Most commonly you will see green, but depending on the type of leaf you may also see purple, red, yellow, or orange.

What about Purple Leaves?

First a bit of science behind purple leaves. Some plants have a lot of compounds called anthrocyanins which can be red or purple in color. If a plant has enough anthrocyanins, the purple will block out the green color even though the chlorophyll is still there. With this particular type of tree we see that in action because the leaves are green in the spring and turn purple for the summer.

During our walk about collecting leaves we came upon a tree with purple leaves. These trees are quite common around here and we were excited to see how it performed in this experiment.

It was interesting to note that the rubbing alcohol turned green initially, then became purple over time.

Another cool fact about purple leaves, the purple colour protects the leaves from sun damage. It’s like tree sunscreen! So cool!

Leaf Coloring Vocabulary Words

Yellow leaves have pigments called xanthophyll .

Carotenoids are the pigment in orange leaves.

And, as we just learned, anthocyanins , give leaves intense red and purple colouring.

Plus of course we have chlorophyll which gives leaves their green colour.

When looking at your chromatography, each colour in your papers is created by those molecules.

What Makes Leaves Change Color?

OK, time for the big question… what makes leaves change colour ? As we know, Chlorophyll gives leaves their green colour and is so dominant that it hides the other colours in the leaves during the spring and summer. But in the fall, chlorophyll in the leaves breaks down, finally allowing the it’s other natural colours to have their moment in the sun!

The beautiful reds, yellows, and oranges might be short lived before the leaves fall to the ground. This year, think about the amazing science behind those changing colours as the trees around you put on their colourful fall displays.

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Science Experiment: Why Do Leaves Change Color?

September 6, 2023 by Sarah 71 Comments

Science experiments for kids are big in our home. My kids love them—and so do I. While teaching Kindergarten, I made science experiments a priority. Kids are so naturally curious about the world around them, and science experiments bring this learning fully to life!

This is my ABSOLUTE favorite science experiment of all time—oh yes, I promise it is THAT GOOD!

I even included this science experiment in Kindergarten at Home , my curriculum for children ages 4 to 6. Kindergarten at Home consists of 18 units, each spanning two weeks. This science experiment is part of the unit all about leaves! Your little one will make leaf art, prepare a fresh salad with you in the kitchen, practice their literacy skills through some fun leaf activities, and so much more.

Take a peek inside! Check out Kindergarten at Home right here:  https://shop.howweelearn.com/pages/kindergarten-at-home

Why Do Leaves Change Color?

When asked by my eldest daughter at age 3 (don’t ask me how many years ago that was!), “Why do leaves change color?” I had no answer.

I looked it up and found it too complicated for a 3-year-old, but I explained the best I could and let it be.

I was in Teacher’s College at the time, and wouldn’t you know, a few weeks later, we learned about—and did—a science experiment for kids about leaves changing color!  

It made things so much clearer that I immediately did the experiment with Madeline. And my future Kindergarten classes. And my future children. Multiple times. Yep, it really is that good!

Before we get started, be sure to download your Free Printable with step-by-step instructions for this fascinating Why Do Leaves Change Color? Science Experiment:

Please keep in mind this is my (limited) understanding, having done the experiment and some research in that class. I’d love to share it with you. So…

The Science Experiment!

Leaves contain chlorophyll. Chlorophyll makes the leaves green and is so dominant that it covers up all the other colors in the leaves. To figure out what color a leaf would be without the dominant chlorophyll color, we can separate the colors by doing this science experiment.

This experiment is very simple, and you likely already have everything you need!

• three leaves from the same tree
• rubbing alcohol
• plastic baggie or plastic wrap
• paper coffee filter
• small bowl or pan
• scissors (optional to cut the coffee filter)
• elastics (optional to hold the plastic wrap in place)
• tape (optional to hold coffee filter in place)

1. Have your child break the leaves into tiny pieces and put them in the jar. We decided to try leaves from two different trees to see how the results compare (being careful to keep each pile of leaves separate).

2. Pour rubbing alcohol over the leaves until they are just covered. A feline supervisor is optional, but highly encouraged.

3. Mash and stir the leaves into the rubbing alcohol until the alcohol turns slightly green. Really give it a good mashing—this is key. We used a plastic spoon so we could really mush and poke the leaves without worrying about accidentally breaking the jar.

4. Cover the jar with a baggie or plastic wrap. We used a rubber band to help hold it in place. Place the jar in a bowl and pour hot water into the bowl.

5. Leave the jar in the hot water for at least 30 minutes, occasionally swishing the jar to stir the leaves. The alcohol should be a dark green (leave longer if needed). If you can resist, wait 45 minutes or even an hour. We found the color difference in the rubbing alcohol at this stage interesting! Is the chlorophyll perhaps stronger in the leaves on the right?

6. Cut a strip from the coffee filter so it can reach the rubbing alcohol. Place it in the jar, and secure it with a small piece of tape at the top edge if needed.

7. The liquid will travel up the coffee filter, and the colors will separate as the alcohol evaporates off the coffee filter. Let this happen for about an hour for the full effect.

There was definitely a difference between the two types of leaves we chose! While it was difficult to capture in the photo, we could see a lot of orange-red tones in the coffee filter strip on the left, and more yellow tones in the one on the right. We also noticed that the size of the orange-red tinge in the strip on the left was larger than the yellow tinge in the strip on the right.

And sure enough, the leaves on the left turn an orange-red shade in the fall, and the leaves on the right turn yellow!

So… Why Do Leaves Change Color?

In this science experiment, we used rubbing alcohol and energy (hot water) to separate the colors. You likely saw green and, depending on your leaf type, maybe red, yellow, or orange.

As we know, chlorophyll gives leaves their green color and is so dominant it hides the other colors in the leaves. But in the fall, chlorophyll in the leaves breaks down, allowing the other colors to finally shine through and show their beautiful reds, yellows, and oranges.

I hope you and your little ones enjoyed this experiment! Science is everywhere, and science experiments for kids are such a wonderful way to build on our children’s natural curiosity about the world.

If you’re ready for an entire year of Kindergarten activities planned for you–easy peasy and full of fun and wonder—check out Kindergarten at Home!

Your Complete 36-Week Curriculum for 4-6 Year Olds

https://shop.howweelearn.com/pages/kindergarten-at-home

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September 8, 2013 at 8:02 pm

This is a great experiment. Not even just for the kids – I find it fascinating too! lol. Pinned it!

September 8, 2013 at 11:38 pm

I think so too! I love activities that are as exciting for me as they are for my wee ones! Thank you for pinning 🙂

October 4, 2018 at 10:27 am

my class is doing this experiment today.

September 15, 2013 at 7:02 am

Ooooh what a great experiment. I’ve struggled to explain this too.

Thanks for linking to The Sunday Showcase. I’ve pinned to our board.

September 15, 2013 at 11:03 am

Yes – The leaves changing can be tricky to explain! Hopefully this experiment helps it to make a bit of sense – and if your wee one is too young to grasp it, it makes for a cool magic trick all the same! Thank you for pinning

September 25, 2013 at 1:01 pm

What an awesome investigation. I saw this experiment in several other posts with comments that it didn’t quite work. I think they didn’t “stew” the solution like you did. I asked my second grader about color change and got the lecture on chlorophyll back, but I know she will enjoy this experiment and seeing this color separation on her own. Thank you so much for sharing it – I will be featuring this post in the next edition of Afterschool! And, by the way, we just did a fun experiment of our own – http://learningwithmouse.blogspot.com/2013/09/mystery-substanceself-led-discovery.html

September 25, 2013 at 1:28 pm

Thanks Natalie! Yes, I have done this experiment many times – worked every one! It is important to really mash the leaves and be patient. I hope it works well for you – please come back to let us know! I will definitely head over to check out your experiment – we are always up for Science ideas!

September 27, 2013 at 6:35 pm

Thanks so much for sharing this great Science experiment over at After School!

September 27, 2013 at 7:27 pm

My pleasure – thank you for hosting!

September 30, 2013 at 1:03 am

Very cool! I think my kids would love doing this project. Thanks for sharing at the After School Party

September 30, 2013 at 1:05 am

Thank you Kelly. Yes, I think it is a super cool experiment too – for grown ups as well 🙂

September 30, 2013 at 3:31 am

This is awesome! I’m adding this to my must-do list. My son would LOVE it! Thanks for sharing this at the After School Linky Party. I’m featuring it this week. Stop by tomorrow and share more of your phenomenal ideas!

October 1, 2013 at 1:13 am

What a compliment – thank you! I would love to stop by 🙂 Hope you enjoy with your son … Gotta love Science!!

October 2, 2013 at 6:26 pm

Thank you so much for sharing this, Sarah! We went on a nature hike when the leaves were starting to change colour and I have been thinking that I needed to find something to explain to the kids why the leaves change colour. This is perfect!

I found your post through the Mom’s Library Link-up.

October 8, 2013 at 1:12 am

What a neat experiment! My daughter loves science so I will have to do this with her:) I am featuring you on my blog for the “Hearts for Home Blog Hop”. The link will be posted on Thursday:)

October 8, 2013 at 1:46 am

Thank you so much for the feature Erika! I love Science too.

November 8, 2019 at 1:00 pm

Thanks my class of children are 2….they don’t understand but they LOVE my science experiments……and I have fun doing it with them

October 20, 2013 at 2:41 pm

It’s really interesting that rubbing alcohol will make the colors in leaves separate. We will have to try that. I would love to have you visit and link up with my Entertaining and Educational Friday blog hop. – http://highhillhomeschool.blogspot.com

October 20, 2013 at 7:28 pm

It is interesting Julie. Rubbing Alcohol breaks down the chlorophyll, imitating what happens in the cold weather. I find Science so fascinating too! I would love to link up to your blog hop on Friday – I will try to stop by this week! Thanks for the invite,

November 2, 2013 at 10:18 pm

What a cool experiment! I am bookmarking this one for when the twins are a little older- they will love it (and so will I)! Thanks for linking up to Discover & Explore this week!

November 4, 2013 at 9:40 pm

It really is cool. Glad you think so too! Thanks for having me over to link.

November 8, 2013 at 7:42 am

This is such an interesting experiment! I had no idea you could do that. I’m featuring you today in a collection of weather experiments for kids. Thanks for linking up to Discover & Explore!

November 8, 2013 at 7:10 pm

Thanks Shaunna! Yes, it is a really neat experiment – I am eager to check out the others in your round up! Thank you for the feature 🙂

November 22, 2014 at 5:36 pm

Wow! Can hardly wai to try!

December 1, 2014 at 8:43 pm

Oh I hope you do give it a try Barb! It is one we do every year … very, very cool!

September 3, 2015 at 6:09 pm

If I remember correctly, I think rubbing alcohol will also separate out the colors from a permanent marker (e.g. a Sharpie pen). Wonder if the color from chlorophyll is somehow similar? Might be amusing to do as a side by side experiment. 🙂

September 3, 2015 at 7:21 pm

Fascinating – I don’t know about this one, but I certainly want to find out! Thank you for taking the time to mention it Gwen!

September 25, 2015 at 8:16 am

I came across your blog earlier this week because my oldest asked why leaves changed colors. So, we did this experiment. They had so much fun! We ended up using the colored alcohol as a form of watercolor paint. Well we tried to, it didn’t work very well haha.

September 30, 2015 at 10:58 pm

So happy you found me here Chelsey! Glad your little ones liked the experiment. What a cool idea about the watercolour paint from the alcohol!

September 30, 2015 at 3:23 pm

wow. i love this science project. next week we are doing leaves for the theme of the week. can’t wait to do this with the children. they are going to love it.

September 30, 2015 at 10:54 pm

I bet you they will love it! And I am glad you sound so eager about it too – that will make them love it even more! It’s awesome – and so are you! Thank you for commenting!

October 14, 2015 at 5:12 pm

Something went wrong with our experiment. We don’t have any color on our coffee filter. I stewed it like you said. Should we take the coffee filter out of our jar and let it dry?

October 14, 2015 at 7:48 pm

Hmmm – rats! Sorry to hear! I don’t think taking it out and letting it dry will make a difference (however you might as well give it a try, perhaps you will e able to see a faint line) . Perhaps try again with more leaves, really really mashed. I promise it really does work! Sorry it didn’t for you – very disappointing!

October 15, 2015 at 10:39 pm

Thanks for your help! I think there is a VERY faint line of yellow. It’s hard to tell. I probably didn’t mash it enough. That’s okay. It was a learning experience for my son to figure out what to do when something doesn’t turn out right.

October 25, 2015 at 8:10 pm

Well I certainly love your positive attitude! I bet it will work for you next time – just mash like crazy!!!

October 19, 2015 at 12:14 pm

Oh I love this so much! I can’t wait to try it with my little girls, thank you for posting! 🙂

October 25, 2015 at 8:07 pm

So glad you are excited! Hope you and your little one have fun!

November 18, 2015 at 5:49 am

Hello. I would really like to do this experiment with some children, the only problem is that in my country we don’t have white coffe filtres. Can you recommend me something else to use instead?

December 1, 2015 at 7:43 pm

Hi Myriam – great question. I am afraid I do not know of an alternate, but perhaps somebody else will – anyone?

February 25, 2016 at 2:19 am

It’s good

September 13, 2018 at 11:15 am

How about paper towels?

October 5, 2018 at 8:20 pm

I’m not sure if this would work as effectively – but if you give it a try, please let me know!!

September 8, 2019 at 4:42 am

IF you know a a science teacher, (high school, college, middle school), you might be able to get lab filter paper, or the real deal, chromatography paper. If your alcohol isn’t dark green, let it sit on the leaves overnight, in the dark, and most all of the pigments will be extracted. Yellow are carotenoids, red would be anthocyanins. Which makes me wonder if you could grate carrots and see those pigments.

April 18, 2016 at 5:54 pm

You should also play with purple cabbage juice. It’s a great pH indicator and you can test loads of household items: baking soda, lemon juice, soda… 🙂

April 18, 2016 at 10:34 pm

Thanks for the suggestion! Very much appreciated!

October 16, 2016 at 8:46 pm

We are doing this for my daughter’s 6th grade science fair project. We tried 3 different leaves from our yard. But only 1 gave us a faint yellowish line. I’m not sure if we didn’t mash it well enough. We tried it again using 2 different types of leaves and only one gave us good results. I don’t know if it’s the type of leaves we used. We live in Central Florida and don’t see leaves cancer by much. We will keep on trying till we get better results. It’s too late to change project ideas. ?

October 19, 2016 at 8:24 pm

Oh No! I hope it works for you. A few tips would be to super, duper mash the leaves, leave the filter as long as possible in the mixture, and use really, really hot water. We are in Canada and get a whole plethora of colours in the Autumn. I am not sure if it makes a difference – I hope it works for you!

October 7, 2020 at 5:18 pm

I’m not sure if the use of the metal lid to the canning jar made a difference but I did not get the results I expected. I plan to try again later tonight with plactic wrap. Any ideas for how to make the colors pop?

July 12, 2017 at 12:14 am

Have you tried this experiment with a leaf already changing color? If it’d work, it’d be neat to compare the filter strip of the green leaf to the changing leaf months later.

July 17, 2017 at 10:14 pm

That’s a neat idea!

August 31, 2017 at 2:23 pm

Thank you for sharing this simple science idea. To be honest I had forgotten why leaves turn color, so it was a nice little reminder and education for me as well. -makeitmindful

October 19, 2017 at 10:41 am

Glad you enjoyed it!

September 23, 2017 at 9:31 am

Just tried this and made some modifications that worked really well, so thought I would share. I let the leaves sit for several hours in the alcohol (about 4 hours). Then I poured the liquid into a clear glass cup and added an equal volume of water. It was cool to see what the liquid looked like in the clear cup (it was brightly pigmented at this point)! For the filter paper strips – after sitting for one hour there were some faint bands, but nothing impressive. I let the filter paper strips sit overnight, and WOW – there were vibrant bands in the morning! I tried hydrangea leaves (green – turn yellow in the fall), and red maple leaves. The red maple leaves were especially beautiful – there was a thick band of purple pigment at the top of the evaporation line and lower band of green pigment. Hope this is helpful! Thanks for posting this fun experiment!

October 18, 2017 at 3:02 pm

Thank you so much for taking the time to add your modifications Abbi!

November 2, 2019 at 5:25 pm

Why did you dilute the leaf-alcohol solution with water? Wouldn’t that dilute the pigments as well?

October 7, 2018 at 12:27 pm

That is wonderful example for Extraction and color chromatography.Thank you very much.

October 14, 2018 at 8:59 pm

So glad you liked the post – thank you for taking the time to let me know!

October 28, 2018 at 12:53 am

Instead of rubbing alcohol can I use spirit or nail polish remover.

October 28, 2018 at 9:32 pm

I’m afraid I am not sure the answer to this question Sonali. Hopefully someone else will and will reply to your message. If you give it a try please let me know if it works. Thank you!

June 6, 2019 at 12:15 pm

Tried this yesterday with 3 kindergarten classes. It didn’t really work, even hours later at the end of the day. I even tried mashing the leaves in a food processor. But then I checked today and there was faint colour lines!! Maybe because I used 50% rubbing alcohol? Anyway, cool experiment.

August 9, 2019 at 9:05 pm

Oh Shucks! I have found that really, really mashing the leaves is key – but clearly you did that with the food processor. Maybe it was the lower concentration of alcohol. Sorry it didn’t work! That is so disappointing.

October 21, 2019 at 10:23 am

Does this experiment make the leaves themselves actually change color? Do the leaves turn yellow or red? Or just the paper gets color on it?

November 16, 2019 at 4:28 pm

Hi Amanda, great question. It just makes the PAPER change colour.

November 4, 2020 at 11:48 am

Hey Sarah! I’ve done this experiment before and loved it. But this year my kiddos have taken us into a leaves and tree study now and all the leaves have already fallen off of trees. Will this experiment work with a dead leaf (one I get from the ground)?? Let me know. Thanks!

November 11, 2020 at 9:59 pm

What an interesting question! I don’t believe it will work with a dead leaf, as the process is complete. But I could be incorrect!

July 12, 2021 at 3:36 pm

What do you recommend I use if I don’t have coffee filters?

July 15, 2021 at 8:06 am

Hmmm … great question. I wonder if just paper towel would work? I am not sure! Perhaps someone else in this community will have insight!

September 29, 2021 at 5:48 pm

The kids loved this experiment!! We got great results! Thanks for sharing!!!

October 25, 2021 at 12:37 pm

So awesome!!!!! Thank you for sharing! xo

[…] A science experiment about why leaves change colour would be a great extension for older preschoolers.  And an art extension making a fall tree would […]

[…] Science Experiment: Why Do Leaves Change Color? […]

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Changing of the Leaves

• 3 green leaves from the same tree
• Rubbing alcohol
• 1 glass jar
• Plastic wrap
• Paper Strip from a coffee filter
• Small bowl with hot water in it

Instructions:

• Break the all the leaves into tiny pieces and put them in the jar.
• Pour rubbing alcohol over the leaves until they are just covered.
• Mash and stir the leaves into the rubbing alcohol until the rubbing alcohol turns slightly green.
• Cover the jar with plastic wrap.
• Put the jar in a bowl of hot water for 50 minutes. Gently swish the jar every 10 minutes to stir up the leaves.
• Take plastic wrap off and place paper coffee filter strip into rubbing alcohol. Make sure just one end of the strip is in the alcohol and the other end is near the top of the jar.
• Let the paper strip sit in jar for an hour. Then make your observations.

VIDEO COMING SOON BUT YOU CAN STILL ENJOY THESE AWESOME EXPERIMENTS!

How it Works:

In this science experiment we used the rubbing alcohol and energy (hot water) to separate the colours. You likely saw green, and depending on your leaf type, maybe red, yellow, or orange. Chlorophyll gives leaves their green colour and is so dominant it hides the other colours in the leaves. But in the fall, chlorophyll in the leaves breaks down allowing the other colours to finally shine through and show their beautiful reds, yellows, and oranges.

Extra Experiments:

• How do the results change if you leave the leaves in the rubbing alcohol for an hour and a half instead of 50 minutes?
• How do the results change if you start with leaves that no longer are green?
• How do the results change if you use needles from an evergreen tree?

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Easy Fall Leaf Science Experiment: Exploring Colors & Chlorophyll with Kids

After exploring and Learning About the Parts of Leaves we decided to learn about leaf color and chlorophyll with an easy science experiment.  I found…

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Before you start this activity the first thing is to go and enjoy the outdoors and gather up some fall leaves. The kids and I feel so lucky to live close to mountains that are ablaze with the beautiful fall colors–one of my favorite times of the year! We went on a little hike, gathered leaves, and other nature treasures and headed home to be scientists.

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Fall Leaf Science Experiment

3.  Observe what color the alcohol is. Come back throughout the day and check to see if the alcohol has changed color. We let our experiment sit overnight and in the morning the colors were even darker but the second night resulted in no change.

4.  Discuss experiment and compare differences between the containers. We recently continued our learning with a STEAM Coffee Filter Art Activity to make Fall Leaves that would be a perfect addition to this activity.

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Leaf Chromatography Experiment

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I love this, when I was a young mom I worked so much just as both my kids do now but finding activities such as this are wonderful. I love these hands on experiments to share with my grandkids. I am a young grandma but It gives me an idea to maybe take the kids to share these experiences with the elderly, they love company.

Great activity for my class! I love science and so does my preschoolers!

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This looks really cool. My son is only 3 so may do this multiple time as he gets older but I know the color changing aspect of the experiment will be most appreciated by him.

Hi Kristen, Thanks so much! I totally agree, it’s great for any age!

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What a great idea! I can't wait for the leaves to start turning here. 🙂 Thank you so much for linking up at the Learn & Play Link Up. Your post is featured this week on Christian Montessori Network!

This is a great idea. I have a high-schooler doing Biology dealing with plants, and I have a fourth grader who watches the Biology videos with his sister. This would be great to do along with what we are already doing now! Thanks for the idea!

This is such a great way to learn about leaves. If I can find some containers, we may just have to do this, even though we finished our leaf unit already. I will be sharing your post with my My Fat'er'hs World Kindergarten group on Facebook , because I am sure there are some moms there that still have to do the leaf unit., and would love to do this. Thank you for stopping by and sharing with Littles Learning Link Up. I hope you can stop by again this week. Have a great day!

What a fun idea! I love the hands-on way of exploring the color of fall leaves! #ThoughtfulSpot

Hi Emma, Thanks so much! My kiddos definitely thought watching the experiment magically change during the day was fun.

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Science Projects > Life Science Projects > Leaf Experiments  

Leaf Experiments

Explore the science of fall with these leaf experiments for elementary students up through high school!

Leaf cells have a special feature: pigment-containing chloroplasts in certain cells that enable them to produce energy and their own food through photosynthesis.

What does that mean? Well, the chloroplasts within a cell contain different pigments, which are what gives a leaf its color.

Green chlorophyll is the most common type of pigment, but there are also xanthophylls (yellow), cartenoids (yellow, orange), and anthocyanins (red).

The chlorophylls usually hide the other pigments, except when autumn comes along and chlorophyll begins to break down. This is why leaves turn different colors in the fall.

Leaf Experiment with Photosynthesis

What is photosynthesis ? Simply put, it’s the capture of light energy to produce food. Light energy from the sun is transmitted through a leaf’s cells to chloroplasts, where chlorophyll and other absorbing pigments serve as receptors to collect the energy. In the process of photosynthesis, carbon dioxide from the air is converted into energy-rich carbon compounds called carbohydrates. As this happens, oxygen is given off into the air, providing the oxygen that we breathe.

You can test the importance of light energy in plant growth by doing a simple leaf experiment.

Use 2-3 small plants. (Bean plants are a good choice, as they sprout quickly.)

You’ll need one to be the control, with normal growing conditions, either outside in sunlight or inside by a bright window. See how light effects growth by covering the other test plants with a paper bag or small box during part of the day.

Try covering one for four hours during the morning, and another for the whole day.

Observe changes to the plants over the course of a week. Which grows the best? What is the result of light-deprivation? What might happen to a leaf with reduced photosynthesis?

Leaf Science Experiment with Color

To find out more about leaf pigments , do this next experiment. First, you’ll need to extract pigments from leaves. Collect several green leaves from different trees, a few from each one. Maples and others that have dramatic color changes in the fall will work best, but you can use any deciduous leaves (from trees that lose their leaves in the winter). Tear each set of different leaves into several pieces and place them in a glass beaker or small drinking glass, then add just enough rubbing alcohol to cover them. (You can cover the containers with foil or plastic wrap to keep the alcohol from evaporating into the air.) Put the containers in a dish of hot tap water for about 30 minutes, until the alcohol turns green as the pigments from the leaves are absorbed into it.

Next, test to find out what colors are really present in a leaf . You’ll need coffee filters, filter paper , or chromatography paper for this part of the experiment. Cut a strip out of the middle of a coffee filter, about one inch wide, for each of the leaf sets that you want to test. Tape one end of the paper to a pencil or stick, and suspend it across the container with the other end just touching the alcohol and pigment mixture. A bit of the mixture will travel slowly up the paper. After about 30-90 minutes you should be able to see the ‘green’ color break up into several different colors as the different pigments begin to separate. You’ll see different shades of green, and perhaps other colors as well.

Which type of leaf had the most colorful pigments? Based on your experiment, which trees’ leaves do you think will turn the brightest and least brightest colors this fall? Try the experiment again with evergreen leaves or needles to compare the results.

Leaf Experiments with a Microscope

Taking a closer look at plant cells.

Learn even more about plants by studying different sections of real leaves. You can make your own microscope slide of a leaf section and view it under high power with a compound microscope to see cell detail. All you need is a fresh leaf specimen (use one without many holes or blemishes), a plain glass microscope slide , slide coverslip , sharp knife, and water.

Before you begin, make sure the leaf is clean and dry. Lay it out flat on your working surface and slice about a 1” section crosswise out of the center using the knife. The cells surrounding the central vein of the leaf are what you will want to look at; so make sure you slice across a section of the vein. Then, starting at one of the short ends of the strip (the edges that you did not cut), tightly roll the leaf section. Carefully make several very thin slices off one end of the roll with a razor blade or knife. This is a ‘cross section’ of the leaf.

Make a wet mount on a plain slide with the inner part of the leaf section facing up (so the inner cells are visible). You can do this by adding a drop or two of water over the leaf section and then covering it with the coverslip. Look at the slide with your microscope’s 10x objective to see the general structure, and higher power to see cell detail. Record your observations on a copy of our free Microscope worksheets.

Click here to download the Microscope Observation worksheet (pdf)

Click here to download the Stereo Microscope Observation worksheet (pdf)

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Leaf chromatography

In association with Nuffield Foundation

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Try this class practical using paper chromatography to separate and investigate the pigments in a leaf

Most leaves are green due to chlorophyll. This substance is important in photosynthesis (the process by which plants make their food). In this experiment, students investigate the different pigments present in a leaf, from chlorophyll to carotenes, using paper chromatography.

The experiment takes about 30 minutes and can be carried out in groups of two or three students.

• Eye protection
• Pestle and mortar
• Chromatography paper
• Beaker, 100 cm 3
• Small capillary tube (see note 1)
• Cut-up leaves, or leaves and scissors (see note 2)
• Propanone (HIGHLY FLAMMABLE, IRRITANT), supplied in a small bottle fitted with a teat pipette (see note 3)

Equipment notes

• The capillary tubing can be ‘home-made’ from lengths of ordinary glass tubing (diameter: 3–4 mm) using a Bunsen burner fitted with a flame-spreading (‘fish-tail’) jet.
• A variety of leaves can be used. Best results are obtained from trees or bushes with dark green leaves, eg holly.
• Preferably use teat pipettes that do not allow squirting, eg those fitted to dropper bottles of universal indicator.

Health, safety and technical notes

• Read our standard health and safety guidance.
• Wear eye protection throughout.
• Propanone, CH 3 COCH 3 (l), (HIGHLY FLAMMABLE, IRRITANT) – see CLEAPSS Hazcard HC085A .  The vapour of propanone is HIGHLY FLAMMABLE. Do not have any source of ignition nearby.
• Finely cut up some leaves and fill a mortar to about 2 cm depth.
• Add a pinch of sand and about six drops of propanone from the teat pipette.
• Grind the mixture with a pestle for at least three minutes.
• On a strip of chromatography paper, draw a pencil line 3 cm from the bottom.
• Use a fine glass tube to put liquid from the leaf extract onto the centre of the line. Keep the spot as small as possible.
• Allow the spot to dry, then add another spot on top. Add five more drops of solution, letting each one dry before putting on the next. The idea is to build up a very concentrated small spot on the paper.
• Attach the paper to the pencil using sellotape so that when placed in the beaker, the paper is just clear of its base.
• Place no more than about 10 cm 3 of propanone in the beaker and hang the paper so it dips in the propanone. Ensure the propanone level is below the spot.

Source: Royal Society of Chemistry

The equipment required for using paper chromatography to separate the different pigments in leaves

• Avoid moving the beaker in any way once the chromatography has started.
• Leave the experiment until the propanone has soaked near to the top, and then remove the paper from the beaker.
• Mark how high the propanone gets on the paper with a pencil and let the chromatogram dry.

Teaching notes

This experiment works very well providing care is taken over preparing the spot on the chromatography paper. It should be as small and as concentrated as possible. Encourage students to be patient and to wait until each application is dry before adding the next.

At least three spots should be obtained, and one of these should be yellow due to carotenes.

The extent to which any particular component moves up the paper is dependent not only on its solubility in propanone but also on its attraction for the cellulose in the chromatography paper. The yellow carotene spot (with a higher RF value) tends to move up the paper the furthest.

More resources

Add context and inspire your learners with our short career videos showing how chemistry is making a difference .

Additional information

This is a resource from the  Practical Chemistry project , developed by the Nuffield Foundation and the Royal Society of Chemistry.

Practical Chemistry activities accompany  Practical Physics  and  Practical Biology .

© Nuffield Foundation and the Royal Society of Chemistry

• 11-14 years
• 14-16 years
• Practical experiments
• Chromatography

Specification

• 2. Develop and use models to describe the nature of matter; demonstrate how they provide a simple way to to account for the conservation of mass, changes of state, physical change, chemical change, mixtures, and their separation.
• Chromatography as a separation technique in which a mobile phase carrying a mixture is caused to move in contact with a selectively absorbent stationary phase.
• 6 Investigate how paper chromatography can be used to separate and tell the difference between coloured substances. Students should calculate Rf values.
• Chromatography involves a stationary phase and a mobile phase. Separation depends on the distribution of substances between the phases.
• The ratio of the distance moved by a compound (centre of spot from origin) to the distance moved by the solvent can be expressed as its Rf value: Rf = (distance moved by substance / distance moved by solvent)
• Mixtures can be separated by physical processes such as filtration, crystallisation, simple distillation, fractional distillation and chromatography. These physical processes do not involve chemical reactions and no new substances are made.
• Recall that chromatography involves a stationary and a mobile phase and that separation depends on the distribution between the phases.
• Interpret chromatograms, including measuring Rf values.
• Suggest chromatographic methods for distinguishing pure from impure substances.
• 12 Investigate how paper chromatography can be used to separate and tell the difference between coloured substances. Students should calculate Rf values.
• 2.11 Investigate the composition of inks using simple distillation and paper chromatography
• 2.9 Describe paper chromatography as the separation of mixtures of soluble substances by running a solvent (mobile phase) through the mixture on the paper (the paper contains the stationary phase), which causes the substances to move at different rates…
• C2.1g describe the techniques of paper and thin layer chromatography
• 2.9 Describe paper chromatography as the separation of mixtures of soluble substances by running a solvent (mobile phase) through the mixture on the paper (the paper contains the stationary phase), which causes the substances to move at different rates o…
• C5.1.4 recall that chromatography involves a stationary and a mobile phase and that separation depends on the distribution between the phases
• 3 Using chromatography to identify mixtures of dyes in a sample of an unknown composition
• C3 Using chromatography to identify mixtures of dyes in a sample of an unknown composition
• 1.9.5 investigate practically how mixtures can be separated using filtration, crystallisation, paper chromatography, simple distillation or fractional distillation (including using fractional distillation in the laboratory to separate miscible liquids…
• 1.9.7 interpret a paper chromatogram including calculating Rf values;
• carry out paper and thin-layer chromatography and measure the Rf values of the components and interpret the chromatograms;

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Science Experiment: The Hidden Colors in a Leaf

Why Do Leaves Change Color in the Fall? 

Leaf Science Experiment Supplies

• Coffee filters
• measuring spoons
• isopropyl alcohol
• Clear jars , glasses or beakers
• a mortar and pestle (optional) 

Leaf Chromatography Experiment

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Leaf Chromatography Experiment – Easy Paper Chromatography

Leaf chromatography is paper chromatography using leaves. Paper chromatography is a separation technique. When applied to leaves, it separates the pigment molecules mostly according to their size. The main pigment molecule in green leaves is chlorophyll, which performs photosynthesis in the plant. Other pigments also occur, such as carotenoids and anthocyanins. When leaves change color in the fall , the amount and type of pigment molecules changes. Leaf chromatography is a fun science project that lets you see these different pigments.

Leaf Chromatography Materials

You only need a few simple materials for the leaf chromatography project:

• Rubbing alcohol (isopropyl alcohol)
• Coffee filters or thick paper towels
• Small clear jars or glasses with lids (or plastic wrap to cover the jars)
• Shallow pan
• Kitchen utensils

You can use any leaves for this project. A single plant leaf contains several pigment molecules, but for the most colors, use a variety of leaves. Or, collect several of each kind of leaf and compare them to each other. Good choices are colorful autumn leaves or chopped spinach.

Perform Paper Chromatography on Leaves

The key steps are breaking open the cells in leaves and extracting the pigment molecule and then separating the pigment using the alcohol and paper.

• Finely chop 2-3 leaves or several small leaves. If available, use a blender to break open the plant cells. The pigment molecules are in the chloroplasts of the cells, which are organelles encased within the plant cell walls. The more you break up the leave, the more pigment you’ll collect.
• Add enough alcohol to just cover the leaves.
• If you have more samples of leaves, repeat this process.
• Cover the container of leaves and alcohol and set it in a shallow pan filled with enough hot tap water to surround and heat the container. You don’t want water getting into your container of leaves.
• Replace the hot water with fresh water as it cools. Swirl the container of leaves around from time to time to aid the pigment extraction into the alcohol. The extraction is ready when the alcohol is deeply colored. The darker its color, the brighter the resulting chromatogram.
• Cut a long strip of coffee filter or sturdy paper towel for each chromatography jar. Paper with an open mesh (like a paper towel) works quickly, but paper with a denser mesh (like a coffee filter) is slower but gives a better pigment separation.
• Place a strip of paper into jar, with one end in the leaf and alcohol mixture and the other end extending upward and out of the jar.
• The alcohol moves via capillary action and evaporation, pulling the pigment molecules along with it. Ultimately, you get bands of color, each containing different pigments. After 30 to 90 minutes (or whenever you achieve pigment separation), remove the paper strips and let them dry.

How Leaf Chromatography Works

Paper chromatography separates pigments in leaf cells on the basis of three criteria:

• Molecule size

Solubility is a measure of how well a pigment molecule dissolves in the sol vent. In this project, the solvent is alcohol . Crushing the leaves breaks open cells so pigments interact with alcohol. Only molecules that are soluble in alcohol migrate with it up the paper.

Assuming a pigment is soluble, the biggest factor in how far it travels up the paper is particle size. Smaller molecules travel further up the paper than larger molecules. Small molecules fit between fibers in the paper more easily than big ones. So, they take a more direct path through the paper and get further in less time. Large molecules slowly work their way through the paper. In the beginning, not much space separates large and small molecules. The paper needs to be long enough that the different-sized molecules have enough time to separate enough to tell them apart.

Paper consists of cellulose, a polysaccharide found in wood, cotton, and other plants. Cellulose is a polar molecule . Polar molecules stick to cellulose and don’t travel very far in paper chromatography. Nonpolar molecules aren’t attracted to cellulose, so they travel further.

Of course, none of this matters if the solvent doesn’t move through the paper. Alcohol moves through paper via capillary action . The adhesive force between the liquid and the paper is greater than the cohesive force of the solvent molecules. So, the alcohol moves, carrying more alcohol and the pigment molecules along with it.

Interpreting the Chromatogram

• The smallest pigment molecules are the ones that traveled the greatest distance. The largest molecules are the ones that traveled the least distance.
• If you compare chromatograms from different jars, you can identify common pigments in their leaves. All things being equal, the lines made by the pigments should be the same distance from the origin as each other. But, usually conditions are not exactly the same, so you compare colors of lines and whether they traveled a short or long distance.
• Try identifying the pigments responsible for the colors.

There are three broad classes of plant pigments: porphyrins, carotenoids, and flavonoids. The main porphyrins are chlorophyll molecules. There are actually multiple forms of chlorophyll, but you can recognize them because they are green. Carotenoids include carotene (yellow or orange), lycopene (orange or red), and xanthophyll (yellow). Flavonoids include flavone and flavonol (both yellow) and anthocyanin (red, purple, or even blue).

Experiment Ideas

• Collect leaves from a single tree or species of tree as they change color in the fall. Compare chromatograms from different colors of leaves. Are the same pigments always present in the leaves? Some plants produce the same pigments, just in differing amounts. Other plants start producing different pigments as the seasons change.
• Compare the pigments in leaves of different kinds of trees.
• Separate leaves according to color and perform leaf chromatography on the different sets. See if you can tell the color of leaves just by looking at the relative amount of different pigments.
• The solvent you use affects the pigments you see. Repeat the experiment using acetone (nail polish remover) instead of alcohol.
• Block, Richard J.; Durrum, Emmett L.; Zweig, Gunter (1955).  A Manual of Paper Chromatography and Paper Electrophoresis . Elsevier. ISBN 978-1-4832-7680-9.
• Ettre, L.S.; Zlatkis, A. (eds.) (2011). 75 Years of Chromatography: A Historical Dialogue . Elsevier. ISBN 978-0-08-085817-3.
• Gross, J. (1991). Pigments in Vegetables: Chlorophylls and Carotenoids . Van Nostrand Reinhold. ISBN 978-0442006570.
• Haslam, Edwin (2007). “Vegetable tannins – Lessons of a phytochemical lifetime.”  Phytochemistry . 68 (22–24): 2713–21. doi: 10.1016/j.phytochem.2007.09.009
• McMurry, J. (2011). Organic chemistry With Biological Applications (2nd ed.). Belmont, CA: Brooks/Cole. ISBN 9780495391470.

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Teacher and Student Resources 

The Process of Leaf Color Change

Leaves change color during the autumn because the amounts of pigments change as the leaves prepare to fall from the trees. All leaves gradually lose chlorophyll during the growing season, and this loss accelerates before leaf fall. Under optimal conditions this process of chlorophyll loss is very orderly and allows the plants to resorb much of the nitrogen in the structure of the pigment molecule. Carotenoid pigments are also lost from the plastids during aging, but some of them are retained in the plastids after the chlorophyll is removed; this produces autumn leaves with yellow colors. In unusual cases, sometimes in winterberry holly, a fair amount of chlorophyll is left in the leaves when they fall. Such leaves are a pale green in color, or perhaps yellow-green from the mixture of chlorophyll and carotenoids.

Most interesting are leaves that turn red, because this color is the result of the active synthesis of anthocyanin pigments just before the leaves fall from the trees. This is the most common color of autumn leaves; about 70 % of shrubs and trees at the Harvard Forest produce anthocyanins during the senescence of the leaves. In these leaves, the actual shades of red are the consequences of the amounts of anthocyanin, the retention of carotenoids (or even a little chlorophyll). Anthocyanin and chlorophyll produce brownish colors. Anthocyanins and carotenoids produce orange hues. In some plants the color production is quite uniform, as in hobblebush or blueberry. In other plants, leaves vary between individuals (as sugar maples) or even dramatically within an individual (as red maples), or even within a single leaf (red maples).

View Color Change in Red Oak or Witch Hazel

The Sequence of Changes in a Healthy Leaf

Color Change in Red Oak

Color Change in Witch Hazel

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Why do leaves change color in the fall.

Frederic Edwin Church, “Autumn Woods,” ca. 1870, Cooper Hewitt, Smithsonian Design Museum, gift of Louis P. Church

Autumnal leaves in vibrant hues are a beautiful part of the season, but those leaves are also a vital part of keeping trees alive.

Trees that have leaves that change color in fall are deciduous. (Evergreen trees with needles, which stay green to continue the photosynthesis process through the winter, are coniferous.) Deciduous trees usually have large, broad leaves.

Most of the year, these leaves are green because of the chlorophyll they use to absorb energy from sunlight during photosynthesis. The leaves convert the energy into sugars to feed the tree.

As the season changes, temperatures drop and days get shorter. Trees get less direct sunlight, and the chlorophyll in the leaves breaks down.

The lack of chlorophyll reveals yellow and orange pigments that were already in the leaves but masked during the warmer months. Darker red leaves are the result of a chemical change: Sugars that can get trapped in the leaves produce new pigments (called anthocyanins) that weren’t part of the leaf in the growing season. Some trees, like oaks and dogwoods, are likely to produce red leaves.

How much and how fast leaves transform varies by location on the globe. The best colors are produced when the weather is dry, sunny and cool. Places that are cloudy, damp or warm won’t see the same degree of changing color.

Then, of course, the leaves fall. Trees start building a protective seal between leaves and their branches as the weather turns. They take in as many nutrients as possible from the leaves, but leaves wouldn’t survive the winter and would make trees vulnerable to damage if they remained. When the leaves are cut off from the fluid in the branches, they separate and drop to the ground.

Learn more about fall tree changes from the Smithsonian Science Education Center, which brings science to K–12 teachers and students.

Leaf through autumnal art from across the Smithsonian .

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15 Simple Leaf Science Activities for Preschoolers

August 24, 2020 by Selena Robinson Leave a Comment

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As the air gets cooler and the days get shorter, it won’t be long before we start to see leaves fall off the trees in our neighborhood.

Down here int he South, it takes a little longer for us to see evidence of fall. But once it starts, it’s glorious.

Leaves are always one of my favorite early childhood themes. And they’re also wonderful opportunities to explore science with little ones.

Which is why I’m sharing this list of 15 simple leaf science activities for preschoolers !

These are simple (and engaging) leaf-themed science ideas for preschoolers and kindergartners.

And they also include sensory fun, math skills, and more!

See the full list below.

Be sure to check out our fun Fall Leaves Count and Match Game for more preschool learning inspiration!

As leaves fall off the trees, we get so many opportunities to explore science. We’re literally watching it happen right in front of us.

We can learn about leaf colors and how they change, compare types of leaves to learn about tree species, and much, much more.

Since preschoolers love to go outside, this is a wonderful time of year to explore nature with them.

So these simple leaf science ideas make learning about nature even easier!

If you decide to try any of these this year, I’d love to know which ones your students liked the most!

Simple Leaf Activities for Preschoolers

These simple leaf activities for preschoolers are a wonderful addition to a fall early learning unit!

There are hands-on ideas here for STEM construction, science experiments, observation activities, nature learning, and more!

Printable Leaf Identification Flashcards

Take these printable leaf identification flashcards out for a nature walk! Perfect for helping little ones learn the names of common leaf varieties!

Easy Fall Leaf Science Experiment: Exploring Colors & Chlorophyll with Kids -

Show preschoolers how chlorophyll affects the color of leaves with this simple experiment! This is also good for learning about colors.

Preschool Leaf Science Experiment

Explore textures and states of matter with this preschool leaf science experiment. Also includes ideas for sensory play!

Preschool Leaf Science and Math Exploration

Examine how leaf textures change over time with this leaf exploration activity. Use the suggestions to add a few math concepts as well!

Salt Crystal Leaves Science Experiment and Fall STEM Activity for Kids

Make actual leaf art through this crystallization science experiment! Easy enough for little learners!

Science for Kids: Exploring How Water Travels Through Leaves

Watch leaves "eat" in real time with this simple science activity. Good for teaching children about the importance of water too!

Best fall simple science for kids - why do leaves change colour?

Show preschoolers how leaves change color with the seasons! This is a beautiful and easy color experiment for little ones.

Observing Autumn Leaves Preschool Unit

Preserve leaves with preschoolers to see how leaves change after falling off trees. This is also good for exploring why leaves need tree roots to live.

Autumn STEAM: tracing the veins of a leaf – Go Science Kids

Add a few art activities into your fall science unit with this STEAM idea. Show kids the veins in each leaf and then use those as an art activity!

Science Experiment: The Hidden Colors in a Leaf

You only need a few basic supplies to try this chromatography experiment. It's great for showing preschoolers the hidden colors in fall leaves!

Easy Nature STEAM: Leaf Painting for Kids

Explore the patterns and textures of leaves with this simple leaf painting STEAM activity. Make prints and then examine how leaf vein patterns vary!

How to Preserve Leaves - Easy Fall Science For Kids

Collect lots of fall leaves, pinecones, and acorns and see how to keep for a memory book! This idea is perfect for a fall preschool table.

Autumn Leaves STEAM Absorption Art and Free Template -

Make gorgeous leaf art with this absorption activity! And teach preschoolers how water affects paper too!

Exploring Leaf Shapes - Nature Walk Activity

Show preschoolers how to make scientific observations with this leaf shapes examination activity! Little ones can collect leaves and compare them with others.

FALL STEM Challenge for Kids: Building Leaf Sculptures

Add some STEM learning into your fall leaves unit with this challenge. Preschoolers will love turning leaves into structures that stand up!

How to Make a Leaf Garland

One of our favorite leaf science activities is making a leaf garland.

It’s such a simple way to explore how fall leaves change over time and add a little fall decor to the home.

Plus, kids can try their hand at making art patterns and exploring colors as they work.

See how to make a simple DIY Fall Leaf Garland with little ones for yourself!

Looking for more preschool fall learning ideas? Try some of these!

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Preschool STEAM

STEAM Activities for Preschoolers

Leaf Science Experiments for Preschoolers

This is a fun roundup with a wide variety of activities using leaves for STEAM projects. Whether exploring the green leaves of summer, the red and yellow leaves of autumn, or even artificial leaves, you’ll be amazed at all the ideas leaves have generated for STEAM!

Natural materials such as leaves are perfect for STEAM projects. Leaves are colorful, come in all shapes and sizes, and are fun for young scientists to explore.

Don’t feel limited if natural leaves are not available in your area. There are lots of beautiful store-bought leaves at your local dollar store that can be used for many leaf science activities.

Textured Leaf Rubbings that Really Work / Preschool STEAM

Experiment with frozen leaves  / Preschool Powol Packets

Measure and sort leaves  / Inspiration Laboratories

Why do leaves change color?  / The Preschool Toolbox

How does a leaf get water?  / Learn Play Imagine

Explore colors and chlorophyll  / A Little Pinch of Perfect

Investigate leaves with leaf rubbings  / Learn With Play at Home

Match the leaves / School Time Snippets

Falling leaf art  / Fantastic Fun and Learning

Explore how water travels through leaves  / Buggy and Buddy

Observe leaves drying indoors  / Preschool Toolkit

Crunchy leaves science and math  / Teach Preschool

How do leaves breathe?  / KC Edventures

Do basic math with fall leaves  / View From a Step Stool

Trace the veins of the leaves  / Go Science Girls

Pound leaves to transfer / Carrots are Orange

Leaf chromatography  / Playdough to Plato

Explore leaf shapes / To Be a Kid Again

This roundup makes it easy to experiment and discover with colorful leaves. Add some fall STEAM to your daily activities with these fun leaf activities!

Welcome to Preschool STEAM!

I'm Jamie a STEAM educator and certified art instructor with more than 12 years experience.

My passion is to show how you can inspire and encourage your students’ creativity through hands-on learning and problem-solving.

You do not need to be an artist, scientist or mathematician to make learning fun and engaging for your students.

Let's get started STEAMing!

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At Preschool STEAM we help educators create STEAM experiences that encourage children to be curious, to wonder, to be innovators and to learn through hands-on play.

Kitchen Science Experiments to Try at Home

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• Kitchen Science Experiments

Chlorophyll Chromatography

Part of the show the science of farming.

Ingredients

Instructions

Cut a piece of filter paper about 1cm wide and a bit longer than your jar is tall.

Place your leaf over the top of the strip and roll the coin across the strip about 2cm from the bottom.

Now add a little acetone (nail varnish remover) to the bottom of your jar, half a cm is plenty.

Suspend your strip of paper so the bottom end is sticking a few mm into the acetone. you can hold it up by folding the top end of the strip over forming a hook which will rest over the lip of the jar.

Wait a few minutes, with any luck you should see an interesting effect.

Try the experiment with a strongly coloured leaf like a purple one.

- Chromatography is pretty  much a black art, and is very dependent on the exact type of paper you are using and the solvent you are using, so if it doesn't work try a different type of paper.

- To get more pigment into the paper and produce a better result you can add some leaf juice, let the paper dry out, and then add some more - repeat as many times as you have the patience for.

- Don't let the paper touch the side of the jar, or all the solvent will move up through the gap between the paper and the glass bypassing your sample.

- Doing the experiment inside a jar seems to be important - probably because it gets filled with acetone vapour and so reduces the evaporation rate of the acetone.

- The experiment only seems to work well on filter paper of one sort or another - probably because it is designed not to stick to things where as most other forms of paper are designed to stick to inks, or what you spilt on the table.

You should find that the acetone slowly rises up the paper, and when it reaches the green mark it takes some of the colour with it, and seperating out the colours.

You should find that may purple leaves still contain green pigments - otherwise they couldn't photosynthesize.

Explanation

This is a technique called chromatography, it works because when you put the paper into the solvent, the solvent is drawn up into the paper by surface tension. This means that there is a flow of liquid over the paper. Some substances spend more time in the solvent and others spend more time on the paper, so they move at different speeds, and get separated out.

This is a very powerful technique which is widely used by chemists to separate out mixtures, both analytically to find out what is in the mixture and to separate out different products of a reaction. The speed of each substance is very dependent on the solvent and the substrate (in this case filter paper) so it is a case of finding the right combination of the two to separate out the substances you want to separate.

Why was there more than one colour of green?

This particlular set  of substrate and solvent has been optimised to separate out different forms of chlorophyll - the pigment which gives plants their colour and is sued in photosynthesis (apparently the ideal combination is dry acetone moving over icing sugar, but dry acetone is hard to get hold of at home). There are in fact 4 types of chlorophyll each having a different absorption spectra. There are two main types found in plants, both which look green but one bluer and the other yellower, so you see two greens. This means that the plant can absorb a greater range of colours of light, and wastes less of the spectrum.

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Last updated by Linda Kamp on December 10, 2022 • 25 Comments

Chlorophyll Paintings: Incorporating Art in Science

Chlorophyll paintings are a fun and creative way to integrate art and science as students learn about plants, the process of photosynthesis and how a plant makes its own food. In this post I’ll share how to incorporate art into your plant science activities by having your students make leaf rubbings and paint pictures with chlorophyll.

How to Make Chlorophyll Paintings

Chlorophyll, the green pigment found in a plant’s leaves, is a key ingredient in the process of photosynthesis. It also makes great paint!

For this project you will need:

• 8-10 leaves per student
• a white sheet of paper
• a metal spoon

I ask students to bring in a bag of leaves from home. I also bring in a big bag of extras and several bunches of fresh spinach. Spinach leaves provide lots of green chlorophyll while some other types of leaves don’t provide as much. We discovered it’s best not to use leaves with a waxy feel since they are harder to press and don’t release their chlorophyll easily.

To make the paintings:

• Fold a sheet of white paper in half. (We used tree templates from this  plant life cycle unit. )
• Place several leaves between the folded paper.
• Press firmly on the leaves between the pages and rub with a metal spoon. Avoid plastic spoons as they break when pressed as firmly as necessary.

Some of my students began with leaf rubbings, but most quickly discovered they could actually “paint” their pictures by balling up the leaves and using them as paintbrushes. This took less time, released more chlorophyll, and they were better able to control where the pigment was on the page.

Chlorophyll rubbing tree page

I gave each student a tree outline template and they painted in the leaves with their chlorophyll. As students work, encourage them to compare the different leaves they are using and discuss which release the most chlorophyll and which release very little. (We had a few of this kind.) Ask students to talk about why they think the leaves of some plants work better while others do not. What qualities or attributes of the leaf make it better or worse for this project?

Incorporating Art into Science

As teachers, we may view art as the opposite of science. Science is driven by data, while art is an expression of creativity. But they have more in common than you think. Both artists and scientists closely observe the world around them. They question, explore, and discover. Bringing these two worlds together in your classroom can lead to a deeper understanding of science topics and help students develop creative problem solving skills.

Printable tree page

Through creative art projects like these chlorophyll paintings, we can often excite our hardest to engage learners and create a unique experience for students who struggle to retain information. I’m a big fan of incorporating art into all areas in my classroom and hope this post gives you ideas for how you can too.

Click here for a complete plant life cycle unit with lesson plans, labs & teaching Power Point

Be sure to pin it for later so you’ll have it when you plan!

Would you like a FREE set of plant science posters?

Drop your personal email in below and I’ll send them to you! (Your personal email is best. Schools often block outside emails with attachments.)

You might also like these science activities! Click the pictures to read these posts.

For more art ideas to use in your classroom check out my Pinterest boards, Art for Kids and Educational Crafts for Kids .

Happy teaching!

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25 comments.

October 1 at 8:33 pm

This is an awesome site!! Great ideas!!

January 19 at 7:18 pm

Amazing info I got from your blog, Good content about chlorophyll paintings! I am just reaching out to say keep up the good work.

https://www.ppropainting.com.au/

March 13 at 3:57 am

Thank you for such a wonderful project. Can’t wait for warm weather, and leaves! This is something I can do with my grandchildren.

April 13 at 11:12 am

I don’t have any comments.

April 4 at 3:45 pm

What standard does this fulfill? Thank you!

April 6 at 7:07 am

Hi Sydney, I don’t believe there is an exact standard that it aligns to, however, I use it as an art extension during our plant unit after students learn about plant needs(sunlight)and the process of photosynthesis.

March 11 at 9:34 am

Do you have questions the kids can use in groups to talk about the different leaves/functions/chlorophyll. Thank you

March 15 at 6:47 pm

Unfortunately I don’t Marian, but I do have a lab for investigating how the shape of a leaf helps the plant get the proper amount of sunlight it needs. You can find the post HERE .

November 18 at 12:07 pm

I’m going to do this art project with my class and wanted to ask how long the “paint” lasts. We want to display the art work in February and don’t know if making them in December is too early.

May 21 at 9:33 am

Unable to leave email and name for plant posters, kept getting the following message: Reference Error: Promise is undefined.

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Hello Friends

I’m Linda Kamp, a 20 year primary grade teacher with a passion for creating educational materials that excite students and make learning fun! I'm so glad you're here!

Write an experiment to show that chlorophyll is necessary for photosynthesis.

Experiment: take a potted plant with variegated leaves like croton and keep it in a dark region, away from sunlight for 3 days. this will halt photosynthesis and de-starch the plant. then keep the plant facing the sunlight for 6 to 8 hours the plant can now carry out photosynthesis and produce starch. mark the green areas in the leaf and trace them on a sheet of paper. mark the regions as green and yellow. the green areas contain chlorophyll which is absent in the yellow areas. immerse the leaf in boiling alcohol to decolorize it. the leaf slowly loses its green color, which goes into the alcohol. dip this decolorized leaf in iodine solution. now remove the leaf from the iodine solution and rinse it in distilled water. remove the leaf from distilled water and keep it on a petri dish. observation two - color regions are visible in the leaf. they are reddish-brown and blue-black. conclusion it can be concluded that the earlier green parts of the leaf turn blue-black whereas the yellow parts have become reddish-brown. green parts of the leaf possess chlorophyll; hence they carry out photosynthesis and produce starch, which turns blue-black with iodine..

Is chlorophyll necessary for photosynthesis? How do we prove it with an experiment? [5 MARKS]

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How to Remove Chlorophyll From Leaves

How to Separate Ink From Water

Chlorophyll is the green pigment found in plant leaves. Chlorophyll works by absorbing sunlight and turning it in to energy for the plant. Students can better understand chlorophyll by extracting it from leaves. This will show students what the true color of the leaf is, which they can also see in autumn just before the leaves fall off trees.

Pour 2 cups water into a medium-sized pot and set it on the stove. Turn the heat setting to high.

Bring the water to a boil and set your leaf in it for two minutes.

Remove the pot from the heat. Use a slotted spoon to take the leaf out.

Pour 1 cup rubbing alcohol into a tall, heat-safe glass and set it in the middle of the pot of hot water. You don't want the water from the pot to spill into the glass.

Place the leaf in the glass with the rubbing alcohol. Make sure the leaf is entirely covered with the rubbing alcohol.

Wait one hour and return to check on the leaf. The leaf should now show its true color. The rubbing alcohol will now be green. The green in the rubbing alcohol is the removed chlorophyll.

Things You'll Need

• This experiment makes a great science project for children to use in a science fair.
• Children should not boil water. An adult should be present for this part of the experiment.

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• Texas Tech University: Plant Pigment Chromatography
• Linus Pauling Institute: Chlorophyll and Chlorophyllin

About the Author

Alicia Bodine has been a professional writer for 13 years. She has produced thousands of articles for online publications such as Demand Studios, GoBankingRates and WiseGeek. Bodine is passionate about gardening, travel, education and finance. She has received awards for being a top content producer.

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ORIGINAL RESEARCH article

Assessing the efficacy of natural soil biotin on soil quality, microbial diversity, and rhododendron simsii growth for sustainable landscape architecture.

• 1 College of Agriculture and Biotechnology, Zhejiang University, Hangzhou, China
• 2 College of Landscape Architecture, Zhejiang A&F University, Hangzhou, China
• 3 Aupro (Hangzhou) Ecological Industry Operations Co., Ltd., Hangzhou, China

Fertilization significantly influences soil quality and its sustainable use in urban garden maintenance. The widespread application of inorganic fertilizers has raised ecological concerns due to their potential environmental impacts. Organic fertilizers, while beneficial, often have slow effects and are costly. Biofertilizers, with their eco-friendly nature and low carbon footprint, are gaining attention for their multifaceted role in supporting plant growth. Despite the focus on fruit trees, vegetables, and medicinal plants, ornamental plants have been understudied. This study aims to evaluate the efficacy of a novel microbial fertilizer, ‘natural soil biotin’, on Rhododendron plants, specifically the Azalea hybrid ‘Carnation’. The study employed a comparative approach to assess the impact of different fertilization strategies on soil properties, microbial diversity, enzyme activity, plant morphology, and physiological parameters. The application of ‘natural soil biotin’ was compared with the use of inorganic and organic fertilizers. The combined application of ‘natural soil biotin’ was found to effectively enhance soil properties and mitigate the impact of other fertilizers on soil pH. It also improved the relative abundance of beneficial microbial groups such as Proteobacteria, Ascomycota, and Basidiomycota. Furthermore, the mixed application significantly increased the activities of urease and sucrase in Rhododendron plants, which promoted their growth, development, and stress resistance. The results indicate that the mixed application of ‘natural soil biotin’ with inorganic and organic fertilizers not only improved the soil quality but also enhanced the efficiency of fertilizer utilization. This approach led to increased economic and environmental benefits in Rhododendron cultivation. The findings contribute to the foundation for soil improvement and ecological restoration, suggesting that ‘natural soil biotin’ could be a promising alternative or supplement to traditional fertilization methods in sustainable landscape architecture.

Graphical Abstract .

• A novel microbial fertilizer boasts a rich composition of diverse organic matter-degrading enzymes.

• The mixed application of natural soil biotin effectively improved soil properties.

• Natural soil biotin increased the relative abundance of Proteobacteria , Ascomycota , and Basidiomycota in the rhizosphere soil.

• Natural soil biotin improved the growth and ornamental value of Rhododendron simsii .

1 Introduction

Rhododendron simsii , a member of the Rhododendron genus in the Rhododendron family, is recognized for its evergreen or deciduous shrub-like or small tree-like structure. Renowned for its vibrant flower colors, diverse flower types, and graceful appearance, it has long been favored for landscaping and container gardening, particularly in China, where it is considered among the top 10 traditionally famous flowers ( Nie et al., 2024 ). The extensive germplasm resources of Rhododendron have garnered global attention, indicative of its widespread visibility. Beyond its ornamental significance, Rhododendron also has notable economic value, driving extensive research across various disciplines, including cultivation, phytochemistry, taxonomy, reproductive and molecular biology, and the microbial ecology of the rhizosphere ( Basnett and Ganesan, 2022 ; Cheng et al., 2024 ; Duarte et al., 2024 ). Moreover, Rhododendron has a particularly strong demand for fertilization during its peak growth seasons, adhering to the principle of ‘sparse yet frequent application’ to optimize growth and vitality.

Fertilizers serve not only as a nutrient source vital for flower growth but also exert a profound influence on their metabolic processes and ornamental qualities. While conventional chemical inorganic fertilizers offer short-term boosts to agricultural productivity, their inherent limitations cannot be overlooked. These fertilizers pose significant risks of environmental contamination, contributing substantially to carbon emissions. Prolonged reliance on such fertilizers may exacerbate issues like soil compaction and declining soil fertility over time ( Gerke, 2022 ). In comparison, organic fertilizers offer the potential for recycling livestock manure, straw, and other waste materials, thereby enhancing soil conditions. Ideally, they should emerge as the favored option for fertilizer application. However, regulatory oversights in China regarding animal feed have led to the inclusion of excessive heavy metals and antibiotics, resulting in the quality of organic fertilizers being diminished and posing increased risks upon application ( He et al., 2023 ). Moreover, organic fertilizer products are often costly and exhibit slow fertilization efficiency, prompting farmers to favor inorganic alternatives. However, excessive reliance on chemical fertilizers and the application of inadequately fermented or immature organic fertilizers can lead to a sharp elevation in nitrogen (N) or salt levels within the soil, which, in turn, disrupts the soil microbial ecology, rendering the soil infertile and jeopardizes its capacity to sustain plant growth ( Li et al., 2022 ; Zhang C. et al., 2022 ). Challenges regarding soil management not only impact the green development of the ecological environment but also amplify the financial burden associated with conventional organic and inorganic fertilizer applications, thereby impeding the sustainable advancement of landscape architecture. Consequently, the quest for an economical and environmentally friendly alternative fertilizer solution emerges as a pressing imperative, particularly concerning the cultivation of flowers such as azaleas and the sustainable evolution of agricultural production.

Compared to traditional fertilizers, biofertilizers, characterized by their eco-friendly attributes, offer the potential for low input and high output. They not only yield significant economic benefits but also play a pivotal role in environmental preservation. Aligned with the trend of sustainable development, biofertilizers exhibit vast application prospects. Biofertilizers, as an innovative category of biological fertilizers, leverage the metabolic activities of diverse microorganisms to deliver targeted fertilization benefits in agricultural practices, enhancing crop productivity sustainably ( Pu et al., 2022 ). Studies have shown that even the application of general biofertilizers can suppress soil-borne fungal pathogens, bolster beneficial microbial populations, improve soil structure, and support plant survival and the growth of green crops ( Wei et al., 2020 ; Zhang et al., 2020 ). Moreover, biofertilizers capitalize on soil organic matter to enrich the soil with secondary organic compounds and increase total N levels, thereby improving soil fertility ( Wei et al., 2020 ). The external application of these biofertilizers has been demonstrated to boost plant yields by stimulating photosynthesis, sugar content, vegetative growth, and advancing the phenological stages, ultimately enhancing the overall quality of the produce ( Zhao et al., 2018 ; Jia et al., 2020 ). These microorganisms within biofertilizers, each with distinct characteristics, facilitate the transformation of nutrients initially inaccessible to plants in the soil, such as insoluble minerals and organic matter. Through the decomposition activities of these active microorganisms, these nutrients are converted into forms readily absorbed and utilized by plants. Simultaneously, these microorganisms exhibit antagonistic properties against soil-borne pathogens, effectively mitigating the incidence of plant diseases and pests while fostering enhanced nutrient absorption and utilization by plants ( Hou et al., 2022 ; Solanki et al., 2022 ). This study introduces a groundbreaking microbial fertilizer known as ‘natural soil biotin’ derived from the extraction and cultivation of natural soil. Enriched with a spectrum of organic matter-degrading enzymes, this product rapidly decomposes organic substances, fostering a symbiotic relationship within the ecosystem. ‘Natural soil biotin’ significantly enhances the soil’s microbial and nutritional profile, promoting robust root development in garden plants. Notably, ‘natural soil biotin’, is a lab-processed product that is both environmentally friendly and non-hazardous. It is distinguished by its abundance of oligosaccharide metabolites, a diverse array of microorganisms, trace elements, readily available nutrients, and fermentation by-products and secondary metabolites from its fungal consortia. The formula includes a comprehensive range of microbial communities, such as aerobic and anaerobic bacteria, as well as extremophiles that thrive at both high and low temperatures. Demonstrating exceptional environmental adaptability, ‘natural soil biotin’ retains its efficacy and stability across a wide temperature range from 0°C–70°C and under varying dissolved oxygen levels of 0.2 mg·L −1 or higher. This adaptability sets it apart from conventional biofertilizers, ensuring consistent performance in diverse conditions.

Despite the numerous advantages of biofertilizers, certain limitations remain in both research and application. Firstly, further investigation is warranted into the theoretical research on microbial fertilizer mechanisms. While domestic experts and scholars have made notable progress in this domain, additional exploration is necessary to elucidate the alterations in plant root microbial communities’ post-application of biofertilizers and the mechanisms underlying their synergistic interactions with bacterial strains ( Kong and Liu, 2022 ; Wang K. et al., 2023 ). Secondly, previous studies have predominantly concentrated on enhancing the yield and quality of vegetables while paying less attention to ornamental species ( Teng et al., 2024 ). Consequently, the theoretical framework for microbial fertilizer application in ornamental plants remains relatively underdeveloped. Additionally, there is a pressing need to enhance the stability of microbial fertilizer products and to establish refined quality standards. The key technology of microbial fertilizer involves combining and preserving bacterial strains, which imposes higher requirements on production processes and equipment. To ensure microbial fertilizer quality, it’s necessary to specify quantitative indicators of target bacteria in field applications within product standards ( Bebber and Richards, 2022 ). Moreover, public understanding of microbial fertilizer remains incomplete, potentially creating a gap between research and production. Effective microbial fertilizer application requires consideration of various factors, with selection based on crop and soil characteristics, necessitating region-specific application standards and plans ( Hasan et al., 2023 ).

In this study, the focus was on commonly cultivated garden flowers and trees, specifically Rhododendrons , to investigate the effects of a new microbial fertilizer known as ‘natural soil biotin’ on their growth, development, and soil quality. By comparing different fertilization treatments and analyzing their impacts on the physicochemical properties of the cultivation substrate, microbial community diversity, soil enzyme activity, and Rhododendron growth and ornamental value, the aim was to identify the optimal fertilization method. Overall, this research contributes to reducing fertilizer usage in landscape architecture, enhancing economic and ecological benefits, and laying a foundation for environmental improvement and ecological restoration.

2 Materials and methods

2.1 plant materials.

The tested plants comprised one-year-old potted cuttings of Rhododendron Azalea hybrid ‘Carnation’, cultivated under uniform conditions. The 2-gallon pot is filled with a balanced potting mixture, crafted from a 1:1 ratio of peat soil and pine bark, and supplemented with approximately 6 kg of stroma per pot. This mixture boasts the following fundamental physicochemical properties: pH 5.78 ± 0.01, organic matter content 401.7 ± 37.6 g·kg −1 , total N content 9.75 ± 0.14 g·kg −1 , hydrolyzable N content 328.7 ± 27.0 mg·kg −1 , available phosphorus (P) content 556.7 ± 18.5 mg·kg −1 , and available potassium (K) content 212.3 ± 10.3 mg·kg −1 .

The liquid microbial fertilizer ‘natural soil biotin’, comprising various complex microbial communities, humic acids, and enzymes, was supplied by Hangzhou Aupro Ecological Industry Operations Co., Ltd. The inorganic fertilizer, labeled as NPK compound fertilizer 15:15:15 with total nutrients ≥45%, was procured from Xinyangfeng Agricultural Technology Co., Ltd. Additionally, sheep manure organic fertilizer with an organic matter content of ≥40% was obtained from Ulanqab Lvxiang Agricultural Production Materials Co., Ltd.

2.2 Plant growth response to fertilization strategies

A pot experiment was conducted to assess the impact of various fertilization treatments on the growth and quality of Rhododendron simsii plants. The following 8 treatments were conducted: (1) Control: watering with clear water only, devoid of fertilizers; (2) T1: applying soil bioactive ingredients alone, comprising base fertilizers at a rate of 50-fold liquid for moistening the substrate, supplemented with monthly topdressing fertilizers at the same rate, amounting to a total of 500 mL of topdressing per pot; (3) T2: applying inorganic fertilizers alone, comprising base fertilizers at a rate of 4 g, with monthly topdressing fertilizers applied at 2 g during the growth period; (4) T3: Mixed application of inorganic fertilizers and soil bioactive ingredients, utilizing base fertilizers at 4 g of inorganic fertilizers mixed with 50-fold liquid of soil bioactive ingredients for substrate moistening, and monthly topdressing fertilizers at 2 g of inorganic fertilizers mixed with 50-fold liquid of soil bioactive ingredients; (5) T4: Application of reduced inorganic fertilizers mixed with soil bioactive ingredients, with base fertilizers at 4 g of compound fertilizers mixed with 50-fold liquid of soil bioactive ingredients for substrate moistening, and monthly topdressing fertilizers at 1 g of compound fertilizers mixed with 50-fold liquid of soil bioactive ingredients; (6) T5: applying organic fertilizers alone, involving base fertilizers at a rate of 120 g, supplemented with monthly topdressing fertilizers at 30 g; (7) T6: Mixed application of organic fertilizers and soil bioactive ingredients, utilizing base fertilizers at 120 g of organic fertilizers mixed with 50-fold liquid of soil bioactive ingredients for substrate moistening, and monthly topdressing fertilizers at 30 g of organic fertilizers mixed with 50-fold liquid of soil bioactive ingredients; (8) T7: Application of reduced organic fertilizers mixed with soil bioactive ingredients, with base fertilizers at 120 g of organic fertilizers mixed with 50-fold liquid of soil bioactive ingredients for substrate moistening, and monthly topdressing fertilizers at 15 g of organic fertilizers mixed with 50-fold liquid of soil bioactive ingredients.

The experiment took place from March 2022 to March 2023 at Shenlongwan Ecological Park in Haining, Jiaxing, Zhejiang Province, situated in the northern Hangjiahu Plain at 30.49059°N, 120.64219°E. The region experiences a subtropical monsoon climate, characterized by an average annual temperature of 15.9°C ( Figure 1 ) and an average annual precipitation of 1,187 mm. The tested plants were placed in an open-air nursery in the park. During periods of intense summer sunlight and high temperatures, black shading nets were used to protect them. Daily standardized cultivation and management practices were implemented throughout the duration of the experiment.

Figure 1 . Historical monthly temperature trend of Haining City (originated from the network: https://lishi.tianqi.com/haining/202405.html ).

2.3 Collection and determination of soil samples for physicochemical indicators

Twenty-four soil samples were systematically gathered from distinct areas—east, south, west, and north fields—of the study site. Each soil sample was taken randomly from five pots per treatment, with the surface layer meticulously cleared away to access the rhizosphere soil. This was extracted using the root shaking technique, homogenized, and any plant debris was meticulously sifted out. The collected samples from these pots were then uniformly mixed and apportioned into three replicates for each treatment. The samples were bifurcated for different analyses: one set was air-dried at room temperature and stored in sealed bags for the assessment of soil physicochemical properties and nutrient levels. The other set was cryopreserved at −80°C in an ultra-low-temperature freezer for future examination of microbial community diversity and soil enzymatic activities. This methodical approach ensured comprehensive data collection for a thorough analysis.

The physicochemical properties of soil samples from each treatment group were assessed following national standard methods. Soil pH was determined using the suspension potential method with a soil-water ratio of 1:2.5 by using a model pHS-25 acidity meter (Shanghai Precise Instruments., China). Organic matter content was determined using the K dichromate oxidation-external heating method ( Sun et al., 2018 ). Total N and hydrolyzable N were determined using the Kjeldahl method ( Mühlhauser et al., 1987 ). Available P was determined using the sodium bicarbonate-molybdate antimony colorimetric method ( Zhang et al., 2016 ), while available K was determined using the ammonium acetate-flame photometric method ( Zhou et al., 2021 ). The parameters including the content of organic matter, total N, hydrolyzable N, available P, and available K were measured by Standard Sci-Tec Innovation (Qingdao) Pharmaceutical Technology Co., Ltd.

2.4 High-throughput sequencing analysis of rhizosphere soil microbial community structure and diversity

Soil samples from the rhizosphere of each treatment were collected and forwarded to Beijing Biomarker Technologies Co., Ltd. for sequencing analysis. DNA extraction from the rhizosphere soil of Rhododendron plants subjected to different treatments was conducted using the TGuide S96 MagBead Soil Genomic DNA Extraction Kit (Tiangen Biotech (Beijing) Co., Ltd.). Each sample was replicated three times. DNA concentration and purity were assessed using a NanoDrop 2000 UV–Vis Spectrophotometer (Thermo Scientific, Wilmington, United States), and DNA integrity was verified using 1.8% agarose gel electrophoresis.

Primers were designed based on conserved regions, with sequencing adaptors added to their ends. The sequence of bacterial 16S rRNA was specifically amplified using the forward primer 338F (5′-ACTCCTACGGGAGGCAGCA-3′) and the reverse primer 806R (5′-GGACTACHVGGGTWTCTAAT-3′). Additionally, the sequence of the fungal ITS region (internal transcribed spacer) was specifically amplified using the forward primer ITS1F (5′-CTTGGTCATTTAGAG GAAGTAA-3′) and the reverse primer ITS2R (5′-GCTGCGTTCTT CATCGATGC-3′). The PCR system comprised 10 μL total volume, including 5 − 50 ng of DNA, 0.3 μL of each primer (10 μM), 5 μL of KOD FX NeoBuffer, 2 μL of dNTP (2 mM each), 0.2 μL of KOD FX Neo polymerase, and ddH 2 O to make up to 10 μL. The PCR amplification program consisted of pre-denaturation at 95°C for 5 min, denaturation at 95°C for 30 s, annealing at 50°C for 30 s, extension at 72°C for 40 s, with 25 cycles, and final extension at 72°C for 7 min, and preservation at 4°C. Fungal diversity analysis was subsequently conducted on the BMK cloud platform. The raw sequence data was deposited in the National Center for Biotechnology Information (NCBI) Sequence Read Archive (SRA) database. 1 We have included sample-wise details of all SRA submissions in Supplementary material , including the number of reads generated and the Phred score for each sample, for the convenience of readers as Supplementary Tables 1 , 2 and Supplementary Tables 3, 4.

The PCR products were purified, quantified, and standardized to create a sequencing library. This library underwent initial quality control, and only qualified libraries were sequenced using the Illumina NovaSeq 6000 platform. High-throughput sequencing produced raw image data files, which were processed to generate raw sequencing sequences through base calling. Initially, the raw data was filtered using Trimmomatic 0.33 ( Bolger et al., 2014 ), followed by identification and removal of primer sequences using Cutadapt 1.9.1 ( Martin, 2011 ). Subsequently, the dada2 method in QIIME2 2020.6 was employed for noise reduction ( Callahan et al., 2016 ; Bolyen et al., 2019 ), paired-end sequence assembly, and removal of chimera sequences, resulting in the final set of effective data (Non-chimeric Reads).

The sequences were clustered at a 97% similarity level using USEARCH 10.0 ( Edgar, 2013 ), with a default threshold of 0.005% of the total number of sequences used to filter OTUs ( Bokulich et al., 2013 ). Subsequently, taxonomic annotation of the feature sequences was performed using the Silva.138 database as a reference with a naive Bayes classifier ( Edgar, 2013 ). Alpha diversity and Beta diversity analyses were conducted using the QIIME2 software. 2

2.5 Measurement of soil enzyme activity

In order to evaluate the fertility and overall soil quality of the rhizosphere soil, soil samples from the root zone of each treated Rhododendron plant were collected. Superoxide dismutase (SOD) activities were assessed using ultraviolet spectrophotometry ( Yu et al., 2023 ), and urease (UE) activities were determined using indophenol blue colorimetry ( Tavares et al., 2021 ). Additionally, sucrase (SC) activities were measured using 3,5-dinitrosalicylic acid colorimetry ( Ge et al., 2018 ). The parameters including the activity of SOD, UE, and SC were measured by Suzhou Comin Biotechnology Co., Ltd.

2.6 Measurement of plant growth indicators

During the cultivation experiment, plant height, crown width, and ground diameter were measured every 2 months, while growth indicators such as the number and length of new branches were assessed at the end of the growth period. Plant height, canopy width, and ground diameter were measured using a steel tape measure (accuracy, 1 mm), a steel tape measure, and an electronic Vernier caliper (accuracy, 0.01 mm), respectively. Other indicators were quantified through visual inspection. During the flowering period of the Rhododendron , the number of flowers per inflorescence, flower diameter, and flowering period were observed and recorded. The number of flowers per inflorescence was determined through visual inspection. Flower diameter was measured using an electronic Vernier caliper (accuracy 0.01 mm) at the maximum diameter of the fully open flower. The flowering period was calculated as the duration from 30% budburst to the wilting of most flowers and the loss of ornamental value.

2.7 Measurement of physiological indicators on plant leaves

2.7.1 chlorophyll content.

After 1 year of cultivation, leaves from Rhododendron plants in each treatment were collected from various directions, including the east field, south field, west field, and north field. The midrib was excised, and the leaves were washed with distilled water before measuring the chlorophyll content. Chlorophyll extraction was performed using the 50% acetone-ethanol mixture grinding method, and the chlorophyll content of the samples was accurately determined using ultraviolet spectrophotometry as described by Ling and Chang-bin (2018) . After centrifugation at 4,000 rpm for 10 min at 25°C, 2.0 mL of the supernatant was taken for analysis with an acetone-ethanol mixture as the blank. The Shimadzu UV-1800 spectrophotometer (Shimadzu UV-1800, Tokyo, Japan) measured optical densities at 665, and 649 nm to calculate the concentrations of chlorophyll a , b , and total chlorophyll according to the formula:

Chlorophyll a concentration: C a = 13.95 × A 665 -6.88 × A 649 , unit mg‧L −1 .

Chlorophyll b concentration: C b = 24.96 × A 649 -7.32 × A 665 , unit mg‧L −1 .

Total chlorophyll concentration: C t = C a + C b = 6.63 × A 665 + 18.08 × A 649 , unit mg‧L −1 .

The parameters of chlorophyll content were measured by Suzhou Comin Biotechnology Co., Ltd.

2.7.2 Physiological indicators of cold resistance

At 40 days of age, mature and healthy leaves from various aboveground nodes of each treated plant were harvested and stored at −20°C for subsequent determination of soluble sugar and soluble protein content, the levels of which can reflect plant stress, nutrient status, and overall health.

The soluble sugar content of the samples was assessed using the anthraquinone coloring method. Initially, soluble sugar was extracted from the sample and subsequently measured using a commercial kit (Suzhou Keming Bio-Tech Co., Ltd.). The soluble sugar content was calculated using the following formula: Soluble sugar (mg·g −1 fresh weight) = [(ΔA + 0.07) ÷ 4.275 × V1] ÷ (W × V1 ÷ V2) = 2.34 × (ΔA + 0.07) ÷ W. In the equation: V1 represents the volume of sample added, 0.04 mL; V2 represents the volume of extract added, 10 mL; Cpr represents sample protein concentration, mg‧mL −1 ; W represents the fresh weight of the sample, g.

The soluble protein content of the samples was determined using the Coomassie brilliant blue method. Initially, soluble protein was extracted from the sample and subsequently measured using a commercial kit (Suzhou Keming Bio-Tech Co., Ltd.). The soluble protein content was calculated using the following formula: Cpr (mg·g −1 ) = (△A + 0.0007) ÷ 7.1265 × V total ÷ W = 0.1403 × (△A + 0.0007) ÷ W. In the equation, V total is the volume of extract (1 mL), and W represents the mass of the sample (g).

2.8 Statistical analysis

Three biological and technical replicates were used for each experiment, with five plants per repetition. Camera pictures were captured using a Canon EOS 600D camera. Graphs were plotted using Excel 2019 (Microsoft, Redmond, WA, United States) and assembled using Microsoft PowerPoint 2019. The data are expressed as means ± SD, computed using SPSS 29.0 analysis software (IBM, Chicago, IL, United States), and subjected to analysis of variance to ascertain statistical significance. Statistical significance was considered at the 95% confidence level ( p  < 0.05). Community analysis plots for each treatment sample were created using the R programming language tool. Adobe Photoshop 2020 was employed to make adjustments and merge the actual photos.

3.1 Effect of different fertilization treatment combinations on rhizosphere soil physical and chemical properties

To assess the impact of natural soil biotin on soil properties and plant growth, soil pH and nutrient content were examined in Rhododendron cultivation under various fertilization treatment combinations ( Table 1 ). Soil pH in CK and T1 increased by 0.10 and 0.15, respectively, compared to the initial pH value of 5.78, remaining within the optimal pH range for Rhododendron growth. Application of inorganic fertilizers, including T2, T3, and T4 led to a significant decrease in soil pH, whereas application of organic fertilizers, including T5, T6 and T7 resulted in a significant increase in soil pH. In comparison with T2 (received inorganic fertilizer), T3 (received inorganic fertilizer and mixed natural soil biotin) demonstrated an increase by 1.03-fold, while T4 (received reduced inorganic fertilizer and mixed natural soil biotin) had its pH increase by 1.04-fold. Conversely, compared to T5 (received organic fertilizer), T6 (received organic fertilizer and mixed natural soil biotin) exhibited a pH decrease by 1.0-fold, while T7 (received reduced organic fertilizer and mixed natural soil biotin) showed a pH decrease by 0.98-fold. Relative to CK, the application of T3 significantly boosted soil organic matter content, available P and K content. While T3 and T4 did not significantly alter soil alkaline N content compared to T2, T6 and T7 did not significantly alter soil alkaline N content compared to T5, they did significantly reduce total N content.

Table 1 . Effects of different fertilization treatment combinations on soil nutrient content.

Fertilizer application significantly influenced the activities of catalase, urease, and sucrase in the soil ( Table 2 ). Organic fertilizers increased catalase and sucrase activities, whereas inorganic fertilizers decreased the activities of catalase, urease, and sucrase. The mixed application of natural soil biotin partially enhanced catalase and sucrase activities. These findings indicate that inorganic fertilizers notably boosted available nutrient content in the soil, while organic and natural soil biotin applications increased soil organic matter and total N content. Moreover, organic fertilizer application enhanced catalase, urease, and sucrase activities, whereas inorganic fertilizer application reduced these activities. However, the mixed application of natural soil biotin improved catalase and sucrase activities.

Table 2 . Effects of different treatments on enzyme activities in rhizosphere soil of Rhododendron.

3.2 Effect of different fertilization treatment combinations on the bacterial community in the rhizosphere soil of Rhododendron

After high-throughput sequencing, a total of 1,921,600 gene sequences were obtained from the 8 sample groups. Following double-ended Reads quality control and splicing, 1,916,441 high-quality sequences were generated, with each sample sequence predominantly ranging between 400 bp and 450 bp in length. The dilution curve, illustrating the rate of new feature appearance with continuous sampling, is depicted in Figure 2A . As sequencing sequences increase, the curve tends to flatten, indicating that species richness in this environment does not continue to rise with increased sequencing quantity. These results suggest that the sample sequences are adequate for subsequent analysis. A Venn plot was employed to analyze differences in the number of bacterial amplification sequence variants (ASVs) in the rhizosphere soil of Rhododendron under different fertilization treatments ( Figure 2B ). The analysis revealed a total of 2,458 ASVs across the 8 samples, with 235 ASVs being characteristic, accounting for 9.6% of the total ASVs and ranging from 18.7 to 21.0% in each treatment group. Compared to the control group, the number of ASVs decreased in the fertilization treatment groups, with the T1 treatment exhibiting significantly fewer characteristic ASVs than CK. Additionally, the T2 treatment showed a significantly lower ASV count than CK. Following the application of inorganic fertilizers, the ASV count increased in T3, while it decreased in T4. A similar trend was observed in comparisons between the CK, T5, T6, and T7 groups.

Figure 2 . Analysis of rhizosphere soil bacterial community diversity under different fertilization treatment combinations. (A) Multi groups rarefaction curves of rhizosphere soil bacterial. (B) Venn analysis of soil bacterial community. T4, Half-supplied inorganic fertilizer + natural soil biotin; T7, half-supplied organic fertilizer+ natural soil biotin; T3, inorganic fertilizer + natural soil biotin; T2, inorganic fertilizer; T1, natural soil biotin; CK, no fertilization; T6, organic fertilizer + natural soil biotin; T5, organic fertilizer.

The Alpha diversity index was employed to assess the species richness and diversity of soil bacterial communities across various fertilization treatments ( Table 3 ). The soil bacterial community richness, as indicated by the Ace and Chao1 indices, spanned 725.33 to 787.71 and 725.33 to 787.67, respectively, with the sequence T5 > T6 > T7 > CK > T3 > T2 > T1 > T4. The application of inorganic fertilizer (T2, T3, T4) and natural soil biotin (T1) decreased bacterial richness, while organic fertilizer (T5, T6, T7) led to an increase. The T3 slightly enhanced richness over the T2, but the T4 saw a lower richness. Organic fertilizer treatments, including T5, T6, and T7, mirrored these findings. Diversity indices, Simpson ranging from 0.9966 to 0.9969 and Shannon from 8.884 to 9.066, followed the pattern T5 > T7 > CK > T6 > T1 > T2 > T3 > T4. The PD_whole_tree index, reflecting community diversity, descended through the treatments in the order of CK, T7, T6, T5, T1, T2, T3, to T4, with all but T7 showing a dip in diversity relative to CK.

Table 3 . Alpha diversity index of soil bacterial community.

The feature sequences of each processed sample were classified and annotated, and the results showed that the species compositions of the treatment groups were similar but had differences in species abundance ( Figure 3A ). Among them, the dominant bacterial phyla in each treatment group included Proteobacteria , Acidobacteria , Gemmatimonadota , Chloroflexi , Actinobacteriota , Myxococcota , and Bacteroidota , accounting for 71.4–78.1% of the total relative abundance. Compared with CK, the relative abundance of Proteobacteria increased in the inorganic fertilizer treatment group (T2) and the mixed inorganic and natural soil biotin treatment group (T3, T4). The relative abundance of Acidobacteria was found to be increased in the organic fertilizer treatment group (T5, T6, and T7), and the results of the bacterial and inorganic fertilizer treatment group (T1, T2, T3, and T4) were similar to those of CK. In addition, the relative abundance of Bacillus subtilis and Pseudomonas aeruginosa decreased in the treatment groups, except for the natural soil biotin treatment group (T1), and the relative abundance of Actinobacteria increased in the treatment groups.

Figure 3 . Analysis of the relationship between soil bacterial community structure and soil physical and chemical properties. (A) Relative abundances of the dominant bacterial phylum. (B) Relative abundances of the dominant bacterial order in soil. (C) PCA analysis between soil bacterial samples. (D) Heatmap of the soil samples. (E) Cladogram of soil bacterial community. The circles radiating from the inside out represent the classification level from phylum to species, and each small circle represents a classification at that level. Its diameter is proportional to relative abundance. Nodes of different colors represent the microbial communities that play an important role in the group represented by that color, while yellow represents no significant difference. (F) LDA value distribution histogram. (G) RDA analysis of bacterial communities and soil physicochemical properties at the genus level. T4, Half-supplied inorganic fertilizer + natural soil biotin; T7, half-supplied organic fertilizer+ natural soil biotin; T3, inorganic fertilizer + natural soil biotin; T2, inorganic fertilizer; T1, natural soil biotin; CK, no fertilization; T6, organic fertilizer + natural soil biotin; T5, organic fertilizer. The length of the arrow represents the impact of environmental factors, and the angle between the angle and the coordinate axis represents the correlation between the environmental factor and the coordinate axis. The smaller the angle, the higher the correlation. The closer the sample point is to the arrow, the stronger the effect of this environmental factor on the sample. The sample is located in the same direction as the arrow, indicating a positive correlation between environmental factors and changes in the sample species community, while the opposite is a negative correlation.

The bacterial orders with higher relative abundance in soil samples from each treatment group comprised Rhizobiales , Vicinamibacterales , Burkholdeiales , Gemmatimonales , Sphingomonadales , SBR1031 , Caulobacterales , and Xanthomonadales ( Figure 3B ). Among them, the Rhizobiales and Vicinamibacterales order was the dominant bacterial order in various treatment groups ( Supplementary Table 2 ). Beta diversity analysis was used to compare the similarity of species diversity among different samples, while Principal Component Analysis (PCA) was used to assess the differences between each group of data on a two-dimensional coordinate map. PC1 and PC2 axes were employed to capture the two feature values reflecting the maximum variance, with distinct treatments represented by different colors. Closer distances between data points indicated higher similarity among processed samples. As depicted in Figure 3C , the 8 sample groups primarily clustered into 3 distinct groups. The bacterial community structures of CK and T1 exhibited notable similarity, differing significantly from other treatments. T2, T3, and T4 demonstrated high similarity, while T5, T6, and T7 exhibited similar patterns. The heatmap, constructed based on a distance algorithm, facilitated the generation of a distance matrix between samples, allowing for an intuitive visualization of differences between two samples through changes in color gradients. Figure 3D illustrates that the results obtained from this approach for analyzing the similarity of bacterial community structures under various fertilization treatments were consistent with the PCA analysis. The Line Discriminant Analysis Effect Size (LEfSe) method was used to compare the bacterial community compositions of each treatment group, and the results showed significant differences ( p  < 0.05) among a total of 36 taxa under the set criteria (LDA > 4; Figures 3E , F ). Redundancy analysis (RDA) was performed on the species diversity at the genus level among the samples to determine the correlation between bacterial communities and soil physicochemical properties ( Figure 3G ). The results showed that pH ( p  = 0.001), available P ( p  = 0.001), and available K ( p  = 0.001) were the main environmental factors affecting the structure of soil bacterial communities. The content of organic matter, total N, and hydrolytic N exhibited a positive correlation with the relative abundance of Xanthomonadales , unclassified A4b, unclassified Bacteria, and other bacterial genera. Conversely, the content of available P and available K was positively correlated with the relative abundance of unclassified Xanthobacteriaceae and unclassified Gemmatimonadaceae but negatively correlated with the relative abundance of unclassified A4b and unclassified Bacteria. The pH value was positively correlated with the relative abundance of unclassified Alphaproteobacteria , unclassified Vicinamidobacteriaceae , Subgroup_10, and other bacterial genera and negatively correlated with the relative abundance of unclassified Xanthobacteriaceae , unclassified SCI_84, and other bacterial genera. These findings suggest that different fertilization combinations exert an influence on the diversity of soil bacterial communities, with the application of natural soil biotin effectively reducing bacterial diversity in the rhizosphere soil. Specifically, the application of inorganic fertilizer alone or in combination with natural soil biotin increased the relative abundance of Proteobacteria , while the application of organic fertilizer increased the relative abundance of Acidobacteria . The dominant bacterial orders in the soil samples were Bacteroidota , Gemmatimonales , Sphingomonadales , SBR1031 , Caulobacterales , and Xanthomonadales .

3.3 Effect of different fertilization treatment combinations on the fungal community in the rhizosphere soil of Rhododendron

A total of 1,349,294 high-quality sequences were generated from 8 sets of samples following double-ended Reads quality control and splicing. At the 97% similarity level, the dilution curves of each treatment flattened as sequencing numbers increased, indicating ample sequencing depth to reliably capture the results ( Figure 4A ). From the 8 groups of samples, 1,454 Amplicon Sequence Variants (ASVs) were identified. Specifically, the ASV counts for CK, T1, T2, T3, T4, T5, T6, and T7 treatments were 680, 710, 648, 661, 576, 579, 668, and 579, respectively, constituting 46.8, 48.8, 44.6, 45.5, 39.6, 39.8, 45.9, and 39.8% of the total ASVs ( Figure 4B ). Among these, 171 ASVs were shared across treatments, representing 11.8% of the total ASV count and 24.1–29.5% of the ASVs in each group. Notably, ASV counts decreased in treatments T1, T2, T3, and T4 compared to CK, except for a significant increase in T1. Treatment T6 exhibited a higher ASV count than T5, while T4 had the same ASV numbers as T5.

Figure 4 . Analysis of rhizosphere soil fungal community diversity under different fertilization treatment combinations. (A) Multi groups rarefaction curves of rhizosphere soil fungal. (B) Venn analysis of soil fungal community. T4, Half-supplied inorganic fertilizer + natural soil biotin; T7, half-supplied organic fertilizer+ natural soil biotin; T3, inorganic fertilizer + natural soil biotin; T2, inorganic fertilizer; T1, natural soil biotin; CK, no fertilization; T6, organic fertilizer + natural soil biotin; T5, organic fertilizer.

As shown in Table 4 , the Ace and Chao1 indices reflecting soil fungi diversity across the 8 treatment groups ranged from 331.33 to 391.37 and 331.33 to 391.33, respectively, displaying the following overall performance trend: T1 > CK > T3 > T2 > T6 > T5 > T4 > T7. Compared with CK, treatment with T1 exhibited an approximately 1.8% increase in fungal community richness, while other treatments showed varying degrees of decrease. Notably, the mixed application of natural soil biotin (T3) slightly increased microbial community richness compared to the only use of inorganic fertilizer (T2), whereas richness was relatively lower in the treatment combining natural soil biotin with a reduction in inorganic fertilizer (T4). Similar trends were observed for organic fertilizer treatments (T5, T6, and T7). Regarding the Simpson and Shannon indices, values ranged from 0.9660 to 0.9890 and from 6.611 to 7.453, respectively, with the overall performance trend as follows: T1 > CK > T6 > T2 > T5 > T7 > T3 > T4. The PD_whole_tree index ranged from 95.12 to 115.64, with the overall performance trend: T1 > CK > T2 > T6 > T5 > T3 > T4 > T7. Except for T1, all treatments exhibited a decrease in diversity compared to CK.

Table 4 . Alpha diversity index of soil fungal community.

The taxonomic annotation results showed that the bacterial community composition structure of each treatment was similar, but the relative abundance of species varied greatly. The dominant bacterial phyla in each treatment included Ascomycota , Basidiomycota , Chytridiomycota , Mortierellomycota , and Olpidiomycota , with relative abundances higher than 10% ( Figure 5A ). The relative abundance of Ascomycota in each treatment group followed the order T6 > T1 > T5 > CK > T7 > T3 > T2 > T4. Notably, compared to CK, treatments involving inorganic fertilizers exhibited a significant decrease in relative abundance. Regarding Basidiomycota , the order of relative abundance was T2 > T7 > T5 > T1 > CK > T6 > T3 > T4, with the highest abundance observed when inorganic fertilizers were applied alone. For the phylum Chlamydomonas , the sequence of relative abundance was T4 > T3 > T2 > CK > T7 > T5 > T6 > T1, with a significant increase in abundance observed with inorganic fertilizer treatment, while a decrease was noted with organic fertilizer treatment. In the case of the Aspergillus phylum , the order of relative abundance was T6 > CK > T2 > T3 > T5 > T7 > T4 > T1, with a decrease observed in other treatments except for a slight increase in T6. Lastly, the relative abundance of the oil pot fungus phylum followed the order T4 > T3 > T7 > CK > T2 > T5 > T6 > T1, with T4 exhibiting notably higher abundance compared to other treatments. The dominant fungal orders in each treatment included Saccharomycetes , Hypocreales , Eurotiales , Pleosporales , Chytridiales , Cladosporales , Agaricales , Sordariomycetidae , and Pezizomycetidae ( Figure 5B ). Relative to CK, the abundance of yeast increased across all treatments, with a rise of 0.6 to 1.6% with inorganic fertilizer application, 3.4 to 4.2% with organic fertilizer, and 1.3% with natural soil biotin. Conversely, Streptomyces abundance decreased with inorganic fertilizer treatments but exhibited an increase in other treatment modalities. The abundance of Pseudomonas aeruginosa decreased in treatments with inorganic fertilizers but increased in those with organic fertilizers. Similarly, Cladosporium abundance decreased with inorganic fertilizer treatments but increased with organic fertilizer treatments. Treatments T5 and T7 exhibited the highest abundance of Agaricales , establishing them as the dominant fungi in these treatments. PCA analysis showed that the fungal community structures of CK, T1, and T6 were similar, while those of T2, T3, and T4 were significantly different ( Figure 5C ). Heatmap analysis corroborated these findings, indicating similarities between some samples from CK, T1, and T6 with those from other treatment groups, consistent with PCA results ( Figure 5D ). Figures 5E and 5F showed significant differences ( p  < 0.05) among a total of 51 taxa under the set criteria (LDA>4). RDA analysis showed that pH, available P, and available K were the main factors affecting the structure of soil fungal communities ( Figure 5G ). Organic matter was the main factor affecting the soil samples of CK and T1 while quick-acting P and K were the main factors affecting the soil samples of T2, T3, and T4. pH was the main factor affecting the soil samples of T5, T6, and T7. The organic matter content was positively correlated with the relative abundance of unclassified Fungi and unclassified Basidiomycota . The content of hydrolyzable N, available P, and available K were positively correlated with the relative abundance of Olpidium and negatively correlated with the relative abundance of Fusarium and unclassified Basidiomycota , and pH value was positively correlated with the relative abundance of Aspergillus , Cladosporium , Candida, and Melanoluca . These results indicate that the application of organic fertilizer and natural soil biotin can increase the number of ASVs and the diversity of the soil fungal community, while the application of inorganic fertilizer can reduce the number of ASVs and the diversity of the soil fungal community.

Figure 5 . Analysis of the relationship between soil fungal community structure and soil physical and chemical properties. (A) Relative abundances of the dominant fungal phylum. (B) Relative abundances of the dominant fungal order in soil. (C) PCA analysis between soil fungal samples. (D) Heatmap of the soil samples. (E) Cladogram of soil fungal community. The circles radiating from the inside out represent the classification level from phylum to species, and each small circle represents a classification at that level. Its diameter is proportional to relative abundance. Nodes of different colors represent the microbial communities that play an important role in the group represented by that color, while yellow represents no significant difference. (F) LDA value distribution histogram. (G) RDA analysis of fungal communities and soil physicochemical properties at the genus level. T4, Half-supplied inorganic fertilizer + natural soil biotin; T7, half-supplied organic fertilizer+ natural soil biotin; T3, inorganic fertilizer + natural soil biotin; T2, inorganic fertilizer; T1, natural soil biotin; CK, no fertilization; T6, organic fertilizer + natural soil biotin; T5, organic fertilizer. The length of the arrow represents the impact of environmental factors, and the angle between the angle and the coordinate axis represents the correlation between the environmental factor and the coordinate axis. The smaller the angle, the higher the correlation. The closer the sample point is to the arrow, the stronger the effect of this environmental factor on the sample. The sample is located in the same direction as the arrow, indicating a positive correlation between environmental factors and changes in the sample species community, while the opposite is a negative correlation.

3.4 Effect of different fertilization treatment combinations on the growth of Rhododendron

The effects of different fertilization treatments on the growth of Rhododendron were examined over a 7-month cultivation period from April to October. This period, characterized by cool weather and suitable temperatures, coincides with the flowering phase of the Azalea. Following the flowering period, Azaleas began to produce new branches and foliage, resulting in rapid increases in plant height and width. Subsequently, during summer, the onset of continuous high temperatures and intense sunlight induces a dormancy phase in Rhododendrons , leading to a significant slowdown in growth rate. As temperatures cool down, typically in autumn, the growth rate of Rhododendron plant height and width accelerates once again. Eventually, with the arrival of winter, Rhododendrons enter another dormant period. Therefore, various growth indicators of Rhododendron were assessed during this period.

The plant height and width of Rhododendron in each treatment group are shown in Figures 6A , B . The plant height followed the order of T3 > T2 > T4 > T1 > T5 > T6 > T7 > CK, with a variation ranging between 43.07 cm and 48.71 cm. Compared with CK, the application of inorganic fertilizers (T2, T3, and T4) significantly increased plant height, while other treatments showed improvements, though the difference was not significant. Among them, the application of natural soil biotin alone (T1) increased plant height by 8.1%, while the application of organic fertilizers (T5, T6, and T7) increased plant height by 3.8 to 6.7%. The height of Rhododendron plants treated with mixed application of inorganic fertilizer and natural soil biotin (T3) was the highest, increasing by 12.5% compared to CK. The plant width followed the order of T3 > T4 > T2 > T5 > T7 > T6 > T1 > CK, with the variations ranging between 48.84 cm to 55.98 cm. Compared with CK, the application of inorganic fertilizers (T2, T3, T4) significantly increased the plant width, while other treatments showed no significant differences. Among them, the single application of natural soil biotin (T1) had the smallest increase of 0.3% in plant width, while the application of organic fertilizers (T5, T6, and T7) increased the plant width by 3.1 to 4.3%. The Rhododendron treated with mixed application of inorganic fertilizer and natural soil biotin (T3) had the highest plant width, an increase of 14.6% compared to CK. As shown in Figures 6C – 6E , the number of new branches showed T3 > T4 > T5 > T2 > T1 > T6 > T7 > CK, while the length of new branches showed T3 > T4 > T2 > T6 > T7 > T5 > T1 > CK. Among all treatments, the mixed application of inorganic fertilizer and natural soil biotin (T3) had the greatest impact on the growth of new branches.

Figure 6 . Effect of different fertilization treatment combinations on the growth of Rhododendron . (A) Plant height. (B) Plant crown diameter. (C) Plant ground diameter. (D) Number of new branches. (E) Length of new branches. (F) Number of flowers per inflorescence. (G) Diameter of a single flower. (H) Florescence. (I) Phenotype of Rhododendron cuttings after one-year trial. (J) Leaf color characteristic phenotype. (K) Chlorophyll content. (L) Soluble protein content. (M) Soluble sugar content. Different lowercase letters indicate significant differences between treatments in the same column ( p  < 0.05). T4, Half-supplied inorganic fertilizer + natural soil biotin; T7, half-supplied organic fertilizer+ natural soil biotin; T3, inorganic fertilizer + natural soil biotin; T2, inorganic fertilizer; T1, natural soil biotin; CK, no fertilization; T6, organic fertilizer + natural soil biotin; T5, organic fertilizer.

The effects of different fertilization treatments on the flowering of Rhododendron are shown in Figures 6F – 6H , with the order being T7 > T4 > T2 > T5 > T6 > CK > T3 > T1. Concerning the size of individual flowers, the order was T3 > T6 > T5 > T4 > T1 > CK > T7 > T2. Notably, the treatment involving the mixed application of inorganic fertilizer and natural soil biotin (T3) exhibited the largest single flower diameter, significantly increasing by 4.2 and 5.2% compared to CK and the only application of inorganic fertilizer (T2), respectively. The results of other fertilization treatments did not significantly differ from CK. In terms of the flowering period duration, Rhododendron displayed the order T7 > T5 > T6 > T1 > T4 > T2 > T3 > CK, with all fertilization treatments extending the flowering period. Compared to CK, the application of inorganic fertilizers (T2, T3, and T4) prolonged the flowering period by 0.8–3.5%, while the sole application of natural soil biotin (T1) increased it by 4.7%, and the application of organic fertilizers (T5, T6, and T7) significantly increased it by 6.5%–9.4%. The longest flowering period (33.92 days) was observed in the treatment involving reduced organic fertilizer and mixed application of natural soil biotin (T7).

The leaves of Rhododendron ‘Carniation’ turned reddish brown in winter. During the experiment, it was found that although each treatment led to a color change, the timing and leaf color varied significantly. As shown in Figure 6J , by the end of November, the leaves of all treatments began to change color, with CK showing the earliest color change and the inorganic fertilizer treatments (T2, T3, and T4) showing the latest color change. By the end of January, the peak of leaf color change was observed, with the leaves of each treatment primarily turning reddish-brown. However, CK, natural soil biotin treatment (T1), and organic fertilizer treatments (T5, T6, and T7) exhibited red leaf coloration, whereas a portion of the leaves within the inorganic fertilizer treatments (T2, T3, and T4) remained dark green. In mid-February, the leaves of Rhododendron began to turn green, with the inorganic fertilizer treatments (T2, T3, and T4) showing the earliest color change, followed by CK and natural soil biotin treatment (T1), and the organic fertilizer treatments (T5, T6, and T7) showed the most delayed change. At this point, the disparity in leaf color between treatments was most apparent. CK’s leaves turned green near the petiole and veins, with reddish-brown edges. The leaves of the natural soil biotin treatment (T1) were similar to those of CK and tended to be yellowish. The leaves of the inorganic fertilizer treatments (T2, T3, and T4) had mostly turned green, with only the edges showing withered yellow or red. The leaves of the organic fertilizer treatments (T5, T6, and T7) were still mainly red in color. The leaves of the single application of organic fertilizer (T5) were yellowish-red with withered leaf tips. The leaves of the mixed application of organic fertilizer and natural soil biotin (T6) were uniformly orange-red in color. The leaves of the reduced application of organic fertilizer and natural soil biotin (T7) had turned green near the petiole, while the leaf tips and edges remained reddish brown. By the end of March, most of the leaves of Rhododendron in each treatment group had turned green. The effects of different fertilization treatments on the chlorophyll content of Rhododendron leaves are presented in Figure 6K . The changing trends of chlorophyll a, chlorophyll b, and total chlorophyll content were generally consistent. The total chlorophyll content ranking for each treatment is as follows: T3 > T2 > T7 > CK > T6 > T4 > T5 > T1. Notably, the mixed application of inorganic fertilizer and natural soil biotin (T3) exhibits the highest chlorophyll content, reaching 0.763 mg·g −1 , representing a significant increase of 13.9% compared to CK. Conversely, the chlorophyll content of the single application of natural soil biotin (T1) was significantly decreased by 10.6% compared to CK, while the total chlorophyll content of other treatments showed no significant deviation from CK. Regarding soluble protein content, the order is as follows: T4 > T6 > T3 > T7 > T1 > T2 > T5 > CK ( Figure 6L ). Although there’s an overall improvement in soluble protein content compared to CK across all fertilization treatments, the differences are not statistically significant. Specifically, the soluble protein content of the single application of natural soil biotin (T1) increases by 0.3% compared to CK (9.42 mg·g −1 ). Compared with a single application of inorganic fertilizer (T2) and organic fertilizer (T5), the soluble protein content increased with the mixed application of natural soil biotin (T3, T6) or the reduced application of respective fertilizers combined with natural soil biotin (T4, T7). Notably, the soluble sugar content in the experimental groups surpassed that of the CK, with the order being T6 > T7 > T6 > T1 > T4 > T2 > T3 > CK ( Figure 6M ). The soluble sugar content of a single application of natural soil biotin (T1), inorganic fertilizer (T2), and organic fertilizer (T5) increased by 12.3, 6.5, and 16.0% compared to CK (60.64 mg·g −1 ), respectively. The highest soluble sugar content was found in using the T6 treatment with mixed application of organic fertilizer and natural soil biotin, which was 70.95 mg·g −1 , a significant increase of 17.0% compared to CK. The soluble sugar content of inorganic fertilizer treatments (T2, T3, and T4) was lower compared to other fertilization treatments. The soluble sugar content of inorganic fertilizer treatments (T2, T3, and T4) was lower compared to other fertilization treatments. Compared with T2 (64.56 mg·g −1 ) using a single application of inorganic fertilizer, treatment T4, involving the reduction of inorganic fertilizer and mixed application of natural soil biotin, exhibited a significant increase in soluble sugar content by 3.5%.

It was found that the mixed application of inorganic and natural soil biotin significantly promoted the growth and flowering of Rhododendron , which could increase the height, width, number, and length of new branches, as well as the number and size of flowers. At the same time, reducing the amount of organic fertilizer and mixing it with natural soil biotin could also prolong the flowering period of Rhododendron . Under different fertilization treatments, the color change process of Rhododendron leaves was found to vary, and there are also differences in chlorophyll content, soluble protein content, and soluble sugar content. Among them, the mixed application of inorganic and natural soil biotin (T3) had a significant promoting effect on the nutrition and color of Rhododendron leaves.

4 Discussion

4.1 the application of natural soil biotin improved soil physicochemical properties and increased soil nutrient contents.

As the foundation of agricultural ecosystems, soil directly impacts the growth and development of plants. Among them, the physical and chemical properties of soil play a crucial role in maintaining soil water and fertilizer capacity, its physical and chemical properties are pivotal in maintaining soil water and fertilizer capacity, nutrient supply, and regulating soil air and heat conditions ( Delgado and Gómez, 2016 ; Kumar et al., 2022 ; Zhang X. et al., 2023 ). Fertilization is an important method for adjusting soil physical and chemical properties, but its effectiveness can significantly vary based on the type, amount, and timing of application. This experiment systematically investigated the impacts of diverse fertilization methods on soil pH and nutrient content through a series of fertilization treatments.

The pH value of soil is a key indicator for measuring soil acidity and alkalinity, directly influencing the availability of nutrients to plants. Optimal plant growth typically occurs within a pH range of 6 to 7, with deviations from this range potentially hindering growth ( Zhang Y. et al., 2022 ). In this experiment, a substrate mixed with peat soil and pine scales was utilized for Rhododendron cultivation, considering the plant’s preference for acidic soil conditions. The initial pH value was 5.78 ± 0.01. By applying different fertilizers, it was observed that inorganic fertilizers notably decreased soil pH, potentially benefiting short-term Rhododendron growth but posing long-term risks such as soil acidification ( Yao et al., 2011 ; Turner et al., 2020 ). Conversely, organic fertilizer application significantly raised soil pH, likely attributed to the production of humic acid during organic fertilizer decomposition, which enhances soil buffering capacity. Notably, the application of natural soil biotin alone had minimal impact on pH, suggesting its efficacy in preserving soil acid–base equilibrium.

In addition to pH value, soil nutrient content is also an important indicator for measuring soil fertility. In this experiment, all fertilization treatments were found to increase the nutrient content of the soil to a certain extent, but there were significant differences among different treatments. The application of natural soil biotin significantly increased soil organic matter and total N content, which was closely related to its function of promoting root absorption and plant growth. However, it is worth noting that the content of hydrolyzable N, available P, and available K did not increase as expected and sometimes even decreased, suggesting that when applying natural soil biotin, there is still a need to combine appropriate amounts of chemical fertilizer to ensure a balanced supply of nutrients ( He, 2022 ). The inorganic fertilizer and organic fertilizer treatment groups showed different characteristics in improving soil nutrient content. Inorganic fertilizer had a particularly significant effect on improving available P and K levels, while the effects with organic fertilizer were relatively mild. When natural soil biotin is mixed with other fertilizers, the organic matter content and available nutrients in the soil are more abundant, which is more conducive to plant absorption and utilization.

Additionally, the focus was on soil enzymes, recognized as vital indicators of soil biological activity and integral to the myriads of biochemical reactions occurring within the soil. The vigor of these enzymes mirrors the soil’s biological activity and is crucial for the circulation of nutrients and energy conversion within the soil ( Wang L. et al., 2023 ). Monitoring shifts in their activity provides valuable insights into the existing fertility levels and the ongoing changes within the soil. In this study, the investigation centered on changes in the activities of catalase, urease, and sucrase. The changes in catalase activity were closely related to soil pH value ( Stpniewska et al., 2009 ), and the effects of different fertilization treatments on its activity showed significant differences. Urease is important for soil N cycling ( Fang et al., 2022 ), and its activity can be influenced by various factors, including soil temperature, humidity, and water and fertilizer management measures. The change in sucrase activity reflects the transformation of organic carbon in the soil ( Zhang Z. et al., 2022 ). The application of organic fertilizer significantly improved sucrase activity, possibly due to the increase in available carbon sources for microorganisms brought about by the addition of rich carbohydrates to the soil, thereby accelerating the rate of soil carbon cycling ( Arnosti et al., 2021 ). In contrast, the application of inorganic fertilizer significantly decreased sucrase activity. However, the treatment that reduced the amount of inorganic fertilizer and mixed it with microbial fertilizer showed significantly higher sucrase enzyme activity than the treatment that applied inorganic fertilizer alone. This may be because urease activity is also influenced by soil temperature, humidity, and water and fertilizer management measures ( Carlos et al., 2022 ; Kalala et al., 2022 ; Ni and Pacholski, 2022 ). Specifically, when the soil temperature is low and the amount of topdressing is large, the conversion rate of soil nitrate N may slow down, leading to accumulation and thus inhibiting urease activity. Soil enzyme activity is intricately connected to the levels and types of organic matter and N in the soil, serving as an indicator of the soil’s nutrient transformation efficiency.

Therefore, fertilization was observed to significantly influence soil physicochemical properties and enzyme activity. Through appropriate combinations of different fertilizers, soil quality and fertility can be effectively enhanced, thereby creating optimal conditions for plant growth. Specifically, the application of natural soil biotin, as a comprehensive soil management approach, not only balanced soil pH and increased nutrient content but also stimulated soil biological activity by modulating enzyme activity, thereby offering robust support for sustainable agricultural development. These findings provide an important basis for a deeper understanding of the impact of fertilization on soil and its relationship with plant growth, as well as theoretical guidance for the scientific and rational formulation of fertilization strategies.

4.2 The application of natural soil biotin reduced the relative abundance and diversity of soil bacterial communities while improving the relative abundance and diversity of fungal communities

As an ecosystem teeming with biodiversity, soil hosts a vast and diverse microbial community, which not only participates in the transformation and cycling of soil nutrients through metabolic activities but also actively promotes plant growth and development, making them of great significance for studying soil fertility status ( Cheng et al., 2022 ; Chaudhary et al., 2023 ; Jiang et al., 2023 ). The rhizosphere, defined as the microenvironment surrounding the root system and heavily influenced by plant roots, serves as a dynamic interface facilitating material exchange between soil and plant ecosystems ( Gregory, 2022 ). Root exudates supply ample carbon and N sources, fostering a microbial population within the rhizosphere that often surpasses that of non-rhizosphere microorganisms. Thus, distinct differences arise in the population structure and diversity of microorganisms inhabiting this zone. These rhizosphere microorganisms play important roles in modulating soil characteristics, thereby influencing plant physiological metabolism and indirectly shaping plant growth and development ( Kong and Liu, 2022 ; Wang D. et al., 2022 ).

This study examined the impact of various fertilization methods on the bacterial and fungal community diversity within the rhizosphere soil of Rhododendron . The results revealed that the single application of natural soil biotin significantly reduced the relative abundance and diversity of soil bacteria while increasing the relative abundance and diversity of fungi. This effect could be attributed to the presence of specific functional microbial communities inherent in natural soil biotin. The application of natural soil biotin potentially stimulated the growth and metabolism of certain dominant microbial populations, consequently diminishing or even eliminating the presence of less advantaged microbial communities ( Hao et al., 2023 ). In contrast, the application of inorganic fertilizer alone resulted in a significant decrease in the relative abundance and diversity of soil bacteria and fungi. This outcome may be attributed to the resultant decrease in soil pH following the application of inorganic fertilizer, which is unfavorable for the survival and proliferation of microorganisms ( Lai et al., 2022 ). In the treatment using mixed application of natural soil biotin and reduced application of inorganic fertilizer, the decrease in soil bacterial and fungal relative abundance and diversity was reduced, indicating that the application of natural soil biotin to some extent alleviated the destructive effect of inorganic fertilizer on soil microbial communities. On the other hand, the application of organic fertilizers resulted in divergent effects on soil microbial communities. While there was a slight increase in bacterial relative abundance and diversity, there was a decrease in fungal relative abundance and diversity. Such variations may stem from the distinct impacts of organic fertilizers on the activity levels of different microbial communities ( Holík et al., 2019 ).

Regarding bacterial community structure, different fertilization methods led to differences in the relative abundance of species in the rhizosphere soil of Rhododendron . Although the main components of soil microbial communities in each treatment were similar, the relative abundances of Proteobacteria , Acidobacteria, and Actinobacteria changed significantly. Among them, the application of inorganic fertilizer significantly increased the relative abundance of Proteobacteria , while the application of organic fertilizer reduced the relative abundance of Acidobacteria . N-fixing nutrient bacteria, including rhizobia within the Proteobacteria phylum , contribute to enhancing soil nutrient content and stimulating plant growth ( Gopalakrishnan et al., 2015 ; Alhammad et al., 2023 ). The Acidobacteria phylum thrives in acidic conditions and may engage in ecological functions such as plant residue degradation and iron cycling ( Ward et al., 2009 ; Wang et al., 2021 ). In terms of fungal community structure, the dominant phyla across all treatments are Ascomycota and Basidiomycota . Ascomycota , known for decomposing recalcitrant organic matter, plays a pivotal role in nutrient cycling ( Beimforde et al., 2014 ; Rathnayaka et al., 2023 ). Basidiomycota contributes to the degradation of plant debris, thereby supporting ecological balance and material cycling ( Manici et al., 2024a , b ). In this experiment, the application of both natural soil biotin and organic fertilizer increased the relative abundance of Ascomycota . Additionally, the mixed application of natural soil biotin significantly increased the relative abundance of Basidiomycota , which is beneficial for organic matter decomposition and nutrient utilization.

In summary, the application of natural soil biotin in the rhizosphere soil of Rhododendron results in reduced richness and diversity of bacterial communities increased richness and diversity of fungal communities, and alterations in bacterial and fungal community structures. Notably, it enhances the relative abundances of Proteobacteria , Ascomycota , and Basidiomycota , which is beneficial for the decomposition of soil organic matter, promoting nutrient cycling, and supporting plant growth. Conversely, the application of inorganic fertilizers can detrimentally affect soil microbiota, whereas the combined use of natural soil biotin and reduced inorganic fertilizers can mitigate this damage. Organic fertilizers exhibit diverse effects on soil bacterial and fungal communities. Nonetheless, each treatment significantly influences the structure of the rhizosphere soil microbial community of Rhododendron , with these effects closely intertwined with changes in soil physicochemical properties.

4.3 Application of natural soil biotin promoted the growth and stress resistance of Rhododendron

In contrast to the emphasis on high yield and quality in field crops, flower cultivation prioritizes ornamental value, stress resistance, and simplified management practices ( Darras, 2020 ). Horticultural researchers have long focused on enhancing flower vigor, prolonging the flowering period, and enhancing flower brightness. Fertilization, a pivotal agronomic practice, profoundly influences the growth and ornamental attributes of flowers ( Kentelky and Szekely-Varga, 2021 ). Nevertheless, improper fertilization methods can result in stunted flower growth, diminished soil quality, and even odor issues.

This study systematically compared the effects of eight different fertilization treatments on the growth, development, and ornamental characteristics of Rhododendron . Parameters such as plant height, width, and diameter are crucial indicators reflecting the growth status of Rhododendron ( Wang et al., 2014 ), collectively forming the plant type, which is pivotal in evaluating flower ornamental value ( Ferrante et al., 2015 ). The experimental results showed that all fertilization treatments promoted Rhododendron plant height and widened plant width, with the mixed application of inorganic fertilizer and natural soil biotin yielding the most pronounced effect. However, under conditions of high temperature and aridity, the group treated with organic fertilizer was notably affected, likely due to heat release during the organic fertilizer decomposition process. Hence, caution is advised when utilizing organic fertilizers in such conditions. In addition to plant type, the number and length of new branches are also important parameters for evaluating the growth status of Rhododendron ( Wang F. et al., 2022 ). In this experiment, all fertilization treatments increased the number and length of new branches to a certain extent, but the difference was not significant. This trend was consistent with the changes in plant height and width, further proving the promoting effects of reasonable fertilization on the growth of Rhododendron .

The ornamental value of flowers is not only reflected in their form but also closely related to the number, size, and flowering period of the flowers. In this experiment, the effects of each treatment on the number of flowers in a single inflorescence of Rhododendron were not significant, but the diameter and flowering period of a single flower were significantly correlated with the fertilization treatment. Notably, the mixed application of inorganic fertilizer and natural soil biotin significantly increased the diameter of a single flower and improved the ornamental appearance of Rhododendron . Moreover, the application of organic fertilizer substantially prolonged the flowering duration, with the treatment involving reduced organic fertilizer and mixed application of natural soil biotin exhibiting the longest flowering period, which effectively extends the ornamental duration of Rhododendron .

Additionally, the study assessed the effects of different fertilization treatments on the color of Rhododendron leaves. Chlorophyll, as a key pigment in photosynthesis, directly affects the photosynthetic efficiency and growth rate of plants ( Zhang Z. et al., 2023 ). The study results indicated that the application of inorganic fertilizer and the combined application of natural soil biotin significantly increased the total chlorophyll content of Rhododendron leaves. In contrast, the sole application of natural soil biotin resulted in a decrease in chlorophyll content. This could be attributed to the close relationship between chlorophyll content and soil nutrient levels, with inorganic fertilizers being capable of rapidly enhancing soil fertility and stimulating chlorophyll synthesis ( Alzamel et al., 2022 ). Although natural soil biotin can promote the absorption of nutrients by plants, it needs to be used in combination with inorganic or organic fertilizers; otherwise, it may lead to insufficient soil nutrients and affect the synthesis of chlorophyll. The control group and the treatment group subjected to bacterial and organic fertilizers exhibited notable leaf discoloration during winter, whereas the treatment group treated with inorganic fertilizers did not manifest this phenomenon, possibly due to the distinct effects of various fertilizers on plant physiological metabolism, though the precise mechanism warrants further investigation. The growth and development of plants are intricately linked to environmental conditions. When environmental changes surpass plants’ tolerance thresholds, stress ensues ( El Rasafi et al., 2022 ), posing significant risks to plant health and potentially resulting in plant demise under severe circumstances. Low temperatures represent a primary adverse factor affecting plant growth, particularly amid the escalating instability of the global climate system. Anomalies in temperature and extreme weather events intermittently occur, posing considerable challenges to agricultural productivity. After exposure to cold stress, Rhododendron s may exhibit symptoms such as leaf shrinkage, browning, slowed growth, and diminished ornamental value, thereby severely constraining their widespread utilization in landscaping ( Verleysen, 2005 ; Dixon, 2019 ).

The soluble protein and soluble sugar content of mature leaves from winter azaleas were analyzed to assess the impact of different fertilization treatments on azalea cold tolerance. Soluble proteins play an important role in maintaining plant cell osmotic regulation, preventing dehydration and freezing, and serving as vital physiological indicators of plant cold tolerance ( Grgac et al., 2022 ). Previous studies have established a positive correlation between soluble protein content and plant cold tolerance under low temperatures ( Zhang et al., 2015 ). In this study, while the soluble protein content in each fertilization treatment increased compared to the control group, the difference was not statistically significant. Soluble sugars serve as additional intracellular protective substances in plants under low temperatures. Cold conditions stimulate the hydrolysis of macromolecular organic compounds such as proteins and starch, thereby increasing the production of soluble sugars ( Kaplan et al., 2006 ). Accumulation of soluble sugars can regulate cell osmotic potential, consequently enhancing plant cold resistance ( Yooyongwech et al., 2009 ). In this experiment, the soluble sugar content significantly increased in each fertilization treatment. Notably, the treatment involving mixed organic fertilizer and natural soil biotin exhibited the highest soluble sugar content, with a significant increase of 17.0% compared to the control group. The increase in soluble sugar content in the organic fertilizer treatment exceeded that of the natural soil biotin treatment alone and surpassed that of the inorganic fertilizer treatment. This suggests that all fertilization treatments, to some extent, enhanced the cold tolerance of Rhododendron , with organic fertilizer exhibiting superior performance, while natural soil biotin and inorganic fertilizer performed slightly less effectively.

In summary, appropriate fertilization practices exerted a substantial influence on the growth, development, and ornamental attributes of Rhododendron . In practical cultivation, fertilization strategies should be strategized in accordance with soil conditions, climatic variations, and the specific growth requirements of Rhododendrons to optimize cultivation outcomes. Additionally, careful consideration must be given to fertilizer selection and application to prevent issues such as improper fertilization, which can result in plant damage and soil quality deterioration.

5 Conclusion

This study provides significant findings on the effects of diverse fertilization strategies on both Rhododendron growth and soil characteristics. The research meticulously evaluated the impact of single and combined applications of inorganic and organic fertilizers, alongside natural soil biotin, on the cultivation of Rhododendron , a popular ornamental species. Through comprehensive assessments of soil physicochemical properties, rhizosphere microbial dynamics, enzymatic activities, and key plant growth indicators, the study illuminated the intricate relationships between fertilization practices and plant health. Importantly, the study discovered that the application of natural soil biotin induced notable improvements in soil quality and significantly enhanced Rhododendron growth. The use of natural soil biotin was particularly effective in enriching the soil, promoting robust microbial communities, and increasing enzyme activities, which are indicative of soil fertility and plant vitality. The study also highlighted the potential of natural soil biotin as a sustainable alternative to traditional fertilizers, capable of reducing dependency on inorganic and organic fertilizers while maintaining or even improving plant growth metrics. The analysis of physiological parameters, including chlorophyll, soluble protein, and soluble sugar content, further underscored the positive influence of natural soil biotin on Rhododendron ’s physiological status. These insights are crucial for the development of eco-friendly and efficient fertilization strategies tailored to ornamental plant cultivation. In conclusion, this study delivers valuable insights and practical recommendations for the horticultural industry. It advocates for the adoption of natural soil biotin as a means to bolster ornamental plant cultivation while ensuring environmental sustainability and economic viability. The findings lay a solid theoretical groundwork for the judicious use of fertilizers, enriching agricultural practices with a nature-oriented approach that prioritizes ecological harmony alongside productivity.

Data availability statement

The original contributions presented in the study are included in the article/ Supplementary material , further inquiries can be directed to the corresponding authors.

Author contributions

ZT: Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Software, Supervision, Validation, Visualization, Writing – original draft, Writing – review & editing. LC: Conceptualization, Formal analysis, Methodology, Writing – original draft. SL: Data curation, Investigation, Methodology, Project administration, Writing – review & editing. KP: Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Writing – original draft. DL: Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Writing – original draft, Writing – review & editing. ZG: Funding acquisition, Resources, Writing – original draft. YW: Investigation, Methodology, Writing – original draft. LH: Conceptualization, Data curation, Formal analysis, Methodology, Resources, Writing – original draft. YC: Conceptualization, Funding acquisition, Investigation, Methodology, Project administration, Resources, Visualization, Writing – original draft, Writing – review & editing.

The author(s) declare financial support was received for the research, authorship, and/or publication of this article. The authors are grateful for the financial support from the Zhejiang University Youth Scientific Research Innovation Project (2016QNA6016), the Research and development of new technologies and new products for sustainable development of Fubaiju Industry in Macheng City, Hubei Province (KHT20230882).

Conflict of interest

YW and ZG were employed by Aupro (Hangzhou) Ecological Industry Operation Co., Ltd.

The remaining authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Publisher’s note

All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.

Supplementary material

The Supplementary material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmicb.2024.1421647/full#supplementary-material

1. ^ http://www.ncbi.nlm.nih.gov/sra/

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Keywords: natural soil biotin, sustainable landscape architecture, soil quality improvement, Rhododendron simsii , soil microbial diversity, enzyme activity

Citation: Teng Z, Chen L, Li S, Pan K, Liu D, Gu Z, Wang Y, Huang L and Chen Y (2024) Assessing the efficacy of natural soil biotin on soil quality, microbial diversity, and Rhododendron simsii growth for sustainable landscape architecture. Front. Microbiol . 15:1421647. doi: 10.3389/fmicb.2024.1421647

Received: 22 April 2024; Accepted: 19 July 2024; Published: 07 August 2024.

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Copyright © 2024 Teng, Chen, Li, Pan, Liu, Gu, Wang, Huang and Chen. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY) . The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

*Correspondence: Li Huang, [email protected] ; Yunwen Chen, [email protected]

Disclaimer: All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article or claim that may be made by its manufacturer is not guaranteed or endorsed by the publisher.

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• Anshul Shyam   ORCID: orcid.org/0000-0001-6373-0420 1 ,
• Dharam Paul Sharma   ORCID: orcid.org/0000-0003-3747-5117 1 ,
• Naveen Chand Sharma   ORCID: orcid.org/0000-0003-0910-5792 1 ,
• Upender Singh 2 &
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The addition of organic materials to the fertilization regime can be instrumental in reducing the amount of chemical fertilizers use, while maintain optimum nutrient status in the trees. The 2 year field experiment was conducted to evaluate the effect of different nutrient source combination in comparison to conventional recommendation on growth, fruit yield and leaf nutrient status of plum cv. ‘Black Amber’ grown in hills of Himachal Pradesh, India. Treatments included conventional fertilizers rate which are recommended to the farmer (NPK 100 ) compared with 10 treatment combinations with reduced rate of chemical fertilizer (NPK 80 ; NPK 70 ; NPK 60 ), organic manure (Farmyardmanure; Vermicompost) and microbial formulations (Biofertilizers; Jeevamrit). Data revealed that NPK 70  + 10% RDN through vermicompost + biofertilizer (T-6) and NPK 70  + 10% RDN through vermicompost + jeevamrit (T-7) yielded the best results in terms of shoot growth, leaf area, leaf chlorophyll content, photosynthesis rate, stomatal conductance and transpiration rate in both the years. Cumulative fruit yield peaked in T-7 by 6% over conventional rate NPK 100 (T-1); there were no difference in fruit yield between T-6 and T-7. Integrated fertilizers combination registered significant influence on the leaf nutritional status, but not consistent. The deviation from optimum (DOP) index revealed excess of leaf phosphorus and iron content with all the treatments application, while negative for other elements however, T-6 maintained a better nutrient status than other treatments. The treatment combinations T-6 and T-7 can be recommended to the farmers for maintaining nutrient balance, thereby ensuring sustaining crop production in subsequent years.

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The data that support the findings of this study are available from the corresponding author [DPS], upon reasonable request.

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Acknowledgements

The authors are thankful to Department of Fruit Science, Dr Y S Parmar University of Horticulture and Forestry, Solan, Himachal Pradesh, India for supporting this research. The authors wishes to thank Mr. Deepak Singha for allowing the research trial to be conducted in his orchard.

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DPS, NCS, US, NR, AS: Conceptualization; AS: Investigation, Methodology, Formal analysis; DPS, NCS: Supervision; AS: Writing original draft, Statistical analysis; DPS, AS: Review manuscript, Editing.

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Supplementary Material 1 Supplementary File 1: 3 Table (Table 1 Optimum leaf nutrient range of plum as suggested by Leece ( 1975 ) and Cref value used in DOP index.; Table 2 Quantity of chemical fertilizer, organic manure, biofertilizer and jeevamrit applied as per the treatment allocation during 2021 and 2022.

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Shyam, A., Sharma, D.P., Sharma, N.C. et al. Optimizing Nutrient Integration for Enhancing Plum Growth, Fruit Yield and Leaf Nutrient Status. J Soil Sci Plant Nutr (2024). https://doi.org/10.1007/s42729-024-01959-8

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"Right there!" he exclaimed.

For decades, this fertile land on the shores of the Ionian Sea has been mainly known for olives, in addition to watermelon and other crops.

But even this part of Greece that sees more rain than other parts of the country is grappling with the effects of drought.

After the warmest winter on record, Greece also experienced the hottest June and July since reliable data collection began in 1960.

"There is no winter," Adamopoulos, 38, told AFP, adding that his property has not received a drop of rain since March.

"No water, no cultivation," said the farmer, whose trees seem to grow right into the Ionian Sea.

Most of Adamopoulos' income currently comes from iceberg lettuce.

But with increasingly arid seasons in sight, he may soon have to give up on some of his lucrative, yet water-intensive crops, such as watermelon.

Adamopoulos is among a small number of Greek growers turning to tropical fruits—mangoes, avocados, lychees, cherimoya and macadamia nuts—which he says are "more resistant" to the increasingly intense heat in the Mediterranean region.

For now, he only grows a few dozen mango and avocado trees on his 80-hectare (198-acre) estate.

The exotic fruits are adapting so well to their new surroundings that Adamopoulos now plans to plant a further 300 trees. He he said he had already received orders for his first harvests, due later this month.

The initiative is part of a study by Greek state agriculture institute Demeter to determine whether tropical fruits could help address the country's looming drought problem.

Not a miracle solution

Study supervisor Teresa Tzatzani says the point is to "find new ways to face this climate change, and make it work in our favour".

"It is hotter all year round now, and this is good for these crops," she said.

Although avocado already grows on the island of Crete, scientists were unsure whether the tree would adapt to conditions on the Greek mainland.

And while mango trees need very little rainfall, the last two winters have been unusually dry, Tzatzani noted.

This type of innovation is essential to save the sector from future climate disasters, said Antonis Paraskevopoulos, head of agriculture for the local region of Triphylia.

But for now, tropical fruits are not a miracle solution.

The programme currently has only a dozen farmers and around 10 hectares under cultivation.

And while it is not intended to replace staple local products such as olives or oranges, it can act as a complement, said Tzatzani, who plans to extend the experiment to other Greek regions.

Neighbouring countries are experiencing similar problems. In Italy, Sicilian farmers have started producing mangoes, bananas and papayas.

The Intergovernmental Panel on Climate Change (IPCC) estimates that the Mediterranean basin, one of the "hotspots" of climate change, will experience more frequent heat waves and droughts.

A 'bad year'?

Theodoros Dimitrakakis, another Greek farmer taking part in the initiative, estimates that it will take years for tropical fruit production to become profitable in Greece.

Despite his enthusiasm for the experiment, the 34-year-old says he can't afford to devote all his time to it, as his main source of income, olive trees, requires all of his attention.

His village, like many in Greece, is often without water for several hours during the day due to scheduled cuts.

Last year, his olive yield was 60 percent below average, Dimitrakakis said.

Despite being an environmental activist during his university years, Dimitrakakis acknowledges that he only recently realised that climate change would impact him so soon.

He now hopes to convince other local farmers, some of whom prefer to think it's just a "bad year".

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JSmol Viewer

Early monitoring of maize northern leaf blight using vegetation indices and plant traits from multiangle hyperspectral data.

1. Introduction

2. materials and methods, 2.1. study area, 2.2. data acquisition and processing, 2.2.1. maize sample data, 2.2.2. multiangle hyperspectral data, 2.3. feature extraction and selection for early monitoring of mnlb, 2.3.1. vegetation indices, 2.3.2. plants traits extracted from hyperspectral data based on hybrid model, 2.3.3. feature selection for early monitoring of mnlb, 2.4. construction and evaluation of model for mnlb early monitoring, 3.1. evaluation of the ability of spectral feature and plant traits for early monitoring of mnlb, 3.1.1. spectral reflectance and vegetation indices, 3.1.2. pts obtained from the inversion of the hybrid model, 3.2. performance evaluation of the mnlb early monitoring model based on various features using multiangle hyperspectral data, 3.3. contribution of vis and pts in the mnlb early monitoring model, 4. discussion, 4.1. vis and pts for early monitoring of mnlb, 4.2. the model for early monitoring mnlb, 4.3. the impact of angles on the early monitoring model of mnlb, 5. conclusions, supplementary materials, author contributions, institutional review board statement, data availability statement, acknowledgments, conflicts of interest.

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Guo, A.; Huang, W.; Wang, K.; Qian, B.; Cheng, X. Early Monitoring of Maize Northern Leaf Blight Using Vegetation Indices and Plant Traits from Multiangle Hyperspectral Data. Agriculture 2024 , 14 , 1311. https://doi.org/10.3390/agriculture14081311

Guo A, Huang W, Wang K, Qian B, Cheng X. Early Monitoring of Maize Northern Leaf Blight Using Vegetation Indices and Plant Traits from Multiangle Hyperspectral Data. Agriculture . 2024; 14(8):1311. https://doi.org/10.3390/agriculture14081311

Guo, Anting, Wenjiang Huang, Kun Wang, Binxiang Qian, and Xiangzhe Cheng. 2024. "Early Monitoring of Maize Northern Leaf Blight Using Vegetation Indices and Plant Traits from Multiangle Hyperspectral Data" Agriculture 14, no. 8: 1311. https://doi.org/10.3390/agriculture14081311

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